Influence of Hmgb1 on Nucleosome Structure and Estrogen Receptor Binding Affinity to Concensus Estrogen Response Element on Nucleosomal Dna

INFLUENCE OF HMGB1 ON NUCLEOSOME STRUCTURE AND ESTROGEN RECEPTOR BINDING AFFINITY TO CONCENSUS ESTROGEN RESPONSE ELEMENT ON NUCLEOSOMAL DNA

Yaw Acheampong Sarpong

A Dissertation

Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

December 2010

Committee:

Dr William Scovell, Chair/ Faculty Advisor

Dr Carmen Fioravanti, Chair/ Faculty Advisor

Dr George Bullerjahn

Dr Carol Heckman

Dr Gary Silverman Graduate Faculty Representative ii

This dissertation is dedicated to

Akwasi Acheampong, my father.

Thank you.

ACKNOWLEDGEMENTS

I would like to extend my gratitude to my chair/faculty advisors, Dr. William Scovell and

Dr Carmen Fioravanti, for their advice and guidance throughout my dissertation. I thank Dr Gary

Silverman, Dr George Bullerjahn, and Dr Carol Heckman for sitting in as committee members. I would like to thank Dr Ron Peterson for making all the DNAs used in the project. You are

appreciated. I would also like to thank all faculty and staff of the Department of Chemistry and

Biological Sciences for giving the opportunity to pursue a graduate degree and to Dr Brecher especially, for the daily food for thought. My sincere thanks go to my teachers from University of Ghana, Dr Jonathan Adjimani and Dr Laud Okine. Lastly I would like to thank everyone who made this project happen. Thank you.

Jah guidance.

Abstract

Previous work from our lab has shown the high mobility group 1 protein

(HMGB1) facilitates the binding of estrogen receptor (ER) to DNAs that contains

consensus estrogen response element (cERE), consensus half-sites (cHERE), direct

repeats of the cHERE and cEREs with different number of base pairs in the spacer

region. This serves as the basis for a new paradigm for estrogen binding. HMGB1 is a dynamic, ubiquitous, “architectural” protein that binds nonspecifically in the minor groove of DNA and has been shown to enhance transcription factor binding to their target sequences and the transcription of a subset of genes by RNA pol II.

This work extends these findings to the effect of HMGB1 on remodeling of nucleosomes, ER binding to HMGB1-remodeled nucleosomes and accessibility of DNase

I and the restriction enzyme, Ava I, to nucleosomes. Although the binding of ER to cERE within a nucleosome effectively does not occur in comparison to that on free DNA, the presence of HMGB1 facilitates ER binding within the nucleosome. A 161 bp DNA was constructed to include four nucleosome positioning sequences and a single, rotationally phased and translationally positioned cERE at the dyad axis or 20 or 40 bps from the dyad axis. Nucleosomes were reconstituted by the salt dilution procedure of the DNA with histones from chicken erythrocyte (CE) oligonucleosomes, which was then purified by sucrose gradient centrifugation. The cERE is positioned so that the major groove containing the cERE is facing outward for optimum ER binding.

Using electrophoretic mobility shift assay (EMSA), we show that there is no significant ER binding to cERE (at either position) in the nucleosome up to ca. 150 nM

ER. However, in the presence of 400 nM HMGB1, ER binds to either position with a KD v

value ca. 50 nM. This is ca. 25-fold weaker than its binding affinity to free DNA. We

show that the DNase I 10 bp patterns and the Exo III digestion is unchanged by the

presence of 400 nM HMGB1. Increasing levels of HMGB1, however, reduces the

mobility of the nucleosome band and suggests that HMGB1, by a non-enzymatic

mechanism, interacts with the nucleosome to generate a new population of “nucleosomes

of altered conformation”. Thus, HMGB1 provides an alternate, ATP-independent

mechanism by which a subset of transcription factors can gain access to their recognition sites within a nucleosome. It also suggests that HMGB1 may cooperate with ATP-

dependent chromatin remodeling complexes to enhance their activity. The HMGB1-

remodeled nucleosome were isolated on a 5-30% sucrose gradient and characterized. We

show that the HMGB1-remodeled nucleosomes are more accessible to DNase I and

restriction enzyme digestion. The HMGB1-remodeled nucleosomes show additional

DNase I sensitive bands and the pattern resembles that obtained for DNase I on free

DNA. Restriction enzyme digest shows that the HMGB1-remodeled nucleosomes

(N’/N”) are very accessible to Ava I compared to canonical nucleosomes.

Cleavage of the histone tail domains also enhances binding of ER to the

nucleosomes. The KD of ER binding to tailless nucleosome (without the addition of

HMGB1) was 50 nM and was further reduced to 25 nM in the presence of 400 nM

HMGB1. Ava I digest shows that these HMGB1-remodeled tailless nucleosomes (N*) are the most accessible to Ava I compared to the canonical nucleosomes or tailless

nucleosome. The HMGB1-remodeled tailless nucleosomes also show additional DNase I

sensitive bands and the pattern resemble that obtained for DNase I on free DNA. vi

The HMGB1-remodeled nucleosomes (with tails and tailless) were, however unstable in high salt, higher temperature and excess DNA, and revert to canonical nucleosomes. There is however, no evidence of dissociation of free DNA or the core histone from the nucleosome complex. vii

Table of contents

Introduction 1. The nucleosome structure and histone/DNA interactions ……..…………… 1 1.1 Histone tail domains……………………………………..……………… 6 1.2 Tailless histones…………………………………………………………. 7 2. Chromatin and remodeling………………………………………………….. 8 2.1 ATP-dependent chromating remodeling...... 11 2.2 ATP-independent chromatin remodeling ………………………………. 11 3. HMGB1……………………………………………………………………... 12 3.1 Effect of HMGB1 on transcription factor binding……………………… 13 3.2 Effect of HMGB1 in DNA repair and V(D)J recombination…………………..……………………………………….. 15 3.3 Effect of HMGB1 in transcription………………………………………. 15 4. Estrogen receptor……………………………………………………………. 16 5. Nucleosome positioning sequence, translation positioned and rotationally phase nucleosomes...……………………………………….…… 18

Materials and methods

1. Construction of plasmids containing cERE…………………………………… 22 1.1 Construction of DNA 4E0…………………………………………….. 26 1.2 Construction of DNA 2E2.……………………………………………. 26 2. Isolation and purification of HMGB1 & HMGB2.…………………..………… 27 2.1 Isolation of calf thymus nuclei………………….……………..………. 27 2.2 Purification of HMGB proteins ………………………………………. 27 2.3 Characterization and storage of HMGB1 and HMGB2 proteins…...…. 29 3. Estrogen receptor alpha (ERα) ………………………………………………. . 29 4. Antibodies to core histone proteins……………………………………………. 30 5. Preparation of oligonucleosomes…………………………………………….... 31 5.1 Isolation of chicken erythrocytes (CE) nuclei ………………………… 31 5.2 Preparation of H1-free oligonucleosomes………………………..……. 32 5.3 Preparation of tailless oligonucleosomes………………………..…….. 33 6. Sequence of DNA……………………………………………………………… 34 7. Isolation of 161 bp DNA from plasmid ……………………………………….. 37 7.1 Preparation of DNA labeled on one strand ………………………..…... 37 7.2 Labeling of DNA………………………………….……………..…….. 37 7.3 Separation of 161 bp DNA fragment from plasmid DNA………..……. 38 7.4 Elution of 161 bp DNA fragment from polyacrylamide gel…………… 38 8. Measurement of specific activity of DNA by scintillation counter……………. 39 9. Preparation of rotationally phase and translationally positioned nucleosomes and tailless nucleosomes containing cERE ……………………... 40 10. Preparation of sucrose gradient……………………………………………… 41 11. Fractionation of nucleosomes…………………………………………………. 41 viii

12. Preparation of HMGB1-remodeled nucleosomes…………………………….. 42 13. Sample preparation of HMGB1-remodeled nucleosomes…………………….... 42 14. ER binding to nucleosomes……………………………………………………. 44 15. DNase I digestion of nucleosomes and modified nucleosomes……………….. 45 15.1 DNase I 10 bp pattern…………………………………….….…….. 45 15.2 DNase I footprint………………………………………………...... 46 16. Exo III digestion of nucleosomes ………………………………………...... 47 17. Ava I digestion on nucleosomes and modified nucleosomes…………..……… 48 18. Preparation of DNA A/G ladder to define position on DNA …………..……… 48 19. Electrophoretic Mobility Shift Assay (EMSA)……………………….………… 49 19.1 Gel Preparation …………………………………………………… 49 19.2 DNA gel electrophoresis……………………………….…………... 50 19.3 Gel Drying…………………………………………………..……... 51 20. Autoradiogram………………………………………………………………… 51 21. SDS-PAGE of proteins………………………………………………………… 52 21.1 Gel Preparation……………………………………………………. 52 21.2 Sample Preparation for proteins ………………………………….. 53 21.3 SDS-PAGE………………………………………………………… 53 21.4 Gel Staining………………………………………………………. 53 21.5 Gel Preservation…………………………………………………… 54 22. Quantification of radioactivity to determine KD values……………………….. 54

Results

Preparation of HMGB1 and rotationally phased & translationally positioned nucleosomes

1. Isolation and purification of HMGB1 and HMGB2 proteins from calf thymus……………………………………………………..…………………. 57 2. Preparation and Isolation of oligonucleosomes ….……...…………………… 59 2.1 Micrococcal nuclease (MN) digestion of chromatin...……………………. 59 2.2 Sepharose CL-4B gel permeation chromatography of MN digested chromatin…………………………………………………………………… 59 3. Preparation of nucleosomes……………………………………………….…… 63 3.1 Diagram of 2E2, 3E1 and 4E0…………………………………………. 63 3.2 Histone exchange by salt dialysis procedure with MN digested chromatin……………………………………...………………………… 65

Characterization of nucleosomes and DNA accessibility assays

4. Binding of ER to DNA ……………………………….…………………….… 68 5. DNase I footprint on the free DNA…………………………………………….. 71 6. Binding of ER to nucleosomes in the presence of 400 nM HMGB1…...………. 73 7. DNase I footprint of ER/cERE in 2E2 nucleosomes …………………………… 78 8. HMGB1 does not alter the rotational position of DNA within nucleosomes…… 80 9. Effect of increasing levels of HMGB1 on nucleosomes structure……………… 82 10. Effect of increasing HMGB1 levels on ER binding to nucleosomes…………… 84 11. Isolation of HMGB1-remodeled nucleosomes (N’/N”)………………………… 87 12. Determination of the presence of core histones in nucleosomes and N’/ N”…… 89 13. ER binding to N’/ N”……………………………………………………………. 91 14. DNase I digestion of nucleosomes and N’/ N”……………..…………………… 94 15. HMGB1 does not alter the translational position of DNA within nucleosomes … 101 16. Ava I restriction enzyme digest of nucleosomes and N’/ N”………………..... 103 17. Effect of temperature on N’/ N”……………………………...... ……………. 106 18. Effect of increasing salt concentration on N’/ N” ...…………………………. 108 19. Stability of N’/ N” in different buffers ……………………………………….. 110 20. Effect of increasing levels of unlabeled DNA on EMSA mobility of N’/ N”… 113 21. Determine if the continuous presence of HMGB1 is required for the stability of N’/ N”……………………………………………………………… 115

Isolation and characterization is tailless nucleosomes

22. Characteristics and reactions with tailless nucleosomes ………………………. 118 22.1 Characteristics of tailless nucleosomes …………………………………… 120 22.2 Effect of removing histone tails from core histones ……………………… 123 22.3 DNase I footprint of ER/cERE in 2E2 tailless nucleosomes ……………… 126 23. Effect of increasing HMGB1 concentration on tailless nucleosome …………… 128 24. Isolation of HMGB1-remodeled tailless nucleosomes (N*) …………………… 130 x

25. ER binding to N* …………………………………………………………… 132 26. DNase I digestion of tailless nucleosomes and N*…………………………. 135 27. Ava I digestion of tailless nucleosomes and N*…………………………….. 141 28. Effect of temperature on N*……………………………………….………… 146 29. Effect of increasing salt concentration on N*...... …………………… 148 30. Effect of increasing levels of unlabeled DNA on EMSA mobility of N*…… 150 31. N* do not alter the translational position of DNA within nucleosomes……… 152 32. DNase I digestion of tailless nucleosomes in the presence of 400 nM HMGB1………………………………………………………………………. 154 xi

Discussion

1. Characteristics of the rotationally phased and translationally positioned nucleosome………………………………………. ……………….. 156 2. HMBG1 enhances ER binding to cERE within nucleosomes ………………… 158 3. Increasing levels of HMGB1 affect EMSA mobility of nucleosomes………… 161 4. HMGB1 remodels nucleosomes into a complex of lower EMSA mobility …... 162 5. HMGB1 can alter the DNase I 10 bp digestion pattern within a remodeled nucleosome …………………………………………………………………. 165 6. Exonuclease III shows that HMGB1 does not affect the translation of DNA in HMGB1 remodeled nucleosomes ………………………………… 168 7. HMGB1 enhances accessibility of Ava I to HMGB1-remodeled nucleosomes ……………………………………………………………….. 169 8. Characteristics of the rotationally phased and translational positioned tailless nucleosome …………………………………………………………. 171 9. HMBG1 enhances ER binding to cERE within tailless nucleosomes ………. 173 10. HMGB1 remodels tailless nucleosomes into a complex of lower EMSA mobility………………………………………………………………. 173 11. HMGB1 can alter the DNase I 10 digestion pattern within a remodeled tailless nucleosome …………………………………………………..……… 175 12. HMGB1 does not affect the translation of DNA in HMGB1 remodeled tailless nucleosomes ……………………………………………… 176 13. HMGB1 enhances accessibility and cutting activity of Ava I to HMGB1-remodeled nucleosomes …..………………………………………... 177

xii

Table of figures

1. Nucleosome core particle ……………………………………………… 2 2. Schematic diagram of core histones ………………… ………………. 4 3. Histone handshake motif of H2A/H2B and H3/H4 ………………… 5 4. Nucleosome core particle ……………………………………………… 9 5. Domain structure of HMGB1 ………………………………………… 13 6. Schematic representation of the functional domain organization of nuclear receptors …………………………………………………… 19 7. ERα DNA binding domain (DBD) binding to cERE…………………. 20 8. Ava I restriction site in plasmid and nucleosome positioning sequences and strategy in construction of DNAs……………………… 24 9. DNA sequence for 2E2 DNA………………………………………….. 35 10. DNA sequence for 4E0 DNA ….……………………………………… 36 11. An 18% SDS-PAGE of purified HMGB1 & HMGB2………………. … 58 12. Micrococcal nuclease digestion of chicken erythrocyte chromatin…….. 61 13. SDS-PAGE of core histone proteins in oligonucleosomes…………….. 62 14. Schematic diagram of DNA used to prepare nucleosomes…………….. 64 15. Sucrose gradient sedimentation profile showing free DNA and nucleosomes……………………………………………………………. 66 16. Sucrose gradient centrifugation fractions analyzed on polyacrylamide gel …………………………………………………… 67 17. ER binding to 4E0 DNA ……………………………………………… 69 18. Binding profile of ER binding to 4E0 ………………………………… 70 19. DNase I footprint on DNA containing either 2E2 & 4E0 ……………… 72 20. ER binding to 4E0 nucleosomes in the absence (A) and presence of 400 nM HMGB1 (B) ………………………………………………… 74 21. Binding profile of ER binding to 4E0 nucleosomes in the presence of 400 nM HMGB1. …………………………………………………… 75 22. ER binding to 2E2 nucleosomes in the presence of 400 nM HMGB1…. 76 23. Binding profile of ER binding to nucleosomes in the presence of 400 nM HMGB1………………………………………………………. 77 24. DNase I footprint of ER/cERE in 2E2 nucleosome……..…………..… 79 25. DNase I 10 bp profile for nucleosomes (2E2) and nucleosomes treated with 400 nM HMGB1………………………………………….. 81 26. Incubation of nucleosomes with increasing levels of HMGB1………... 83 27. ER binding to nucleosomes in the presence of increasing concentration of HMGB1………………………………………………. 85 28. Binding profile of ER binding to nucleosomes in the presence of increasing HMGB1 concentration ………………………………….. 86 29. Sedimentation profiles of remodeled nucleosomes…………………….. 88 30. Supershift assay to determine the presence of core histones in nucleosomes, and remodeled nucleosomes…………………………… 90 31. ER binding to remodeled nucleosomes in the absence (A) and presence (B) of poly (dI-dC)………………………………………… 92 xiii

32. Binding profile of remodeled nucleosomes in the absence and presence of poly (dI-dC)…………………………………………… 93 33. DNase I of canonical nucleosomes and modified nucleosomes (2E2)……………………………………………………… 96 34. DNase I of canonical nucleosomes and modified nucleosomes (3E1)………………………………………………………. 97 35. DNase I of canonical nucleosomes and modified nucleosomes (4E0)………………………………………………………. 98 36. Comparison of DNase I sensitive sites on nucleosomal DNAs …………. 100 37. Exo III digestion on nucleosomes (4E0), with and without 1600 nM HMGB1………………………………………………………… 102 38. Schematic of DNA showing the general position of cERE and the Ava I restriction recognition sequence and cutting sites in all DNAs…. 104 39. Ava I digestion of 4E0 nucleosomes and remodeled nucleosomes……….. 105 40. Effect of temperature on mobility of nucleosomes and remodeled nucleosomes ……………………………………………………………… 107 41. Effect of increasing NaCl concentration on mobility of remodeled nucleosomes……………………………………………………………….. 109 42. Stability of HMGB1-remodeled nucleosomes in different buffers……….. 112 43. Effect of increasing the levels of unlabeled 161 bp DNA on modified nucleosome……………………………………………………… 114 44. Titration of HMGB1-remodeled nucleosome with anti-HMGB1………… 117 45. SDS-PAGE of core and tailless histones in oligonucleosomes…………… 120 46. Sucrose gradient sedimentation profile showing free DNA and tailless nucleosomes……………………………………………………………….. 121 47. Sucrose gradient centrifugation fractions analyzed on polyacrylamide gel... 122 48. ER binding affinity to tailless nucleosomes (4E0) in the (A) absence of HMGB1 and in the (B) presence of 400 nM HMGB1………………….. 124 49. Binding profile of ER binding to tailless nucleosomes in the absence and in the presence of 400 nM HMGB1…………………………………… 125 50. DNase I footprint of tailless nucleosomes (2E2) in the absence and presence of 400 nM HMGB1………………………………………….. 127 51. EMSA mobility of tailless nucleosomes with increasing levels of HMGB1…………………………………………………………………….. 129 52. Sedimentation profiles of HMGB1-remodeled tailless nucleosomes (4E0)... 131 53. ER binding to remodeled tailless nucleosomes in (A) the absence and (B) presence of poly (dI-dC) (4E0)..…………………………………………… 133 54. Binding profile of remodeled nucleosomes in the absence and presence of poly (dI-dC)……………………………………………………. 134 55. DNase I of tailless nucleosomes and HMGB1-remodeled tailless nucleosomes (2E2)………………………………………………………… 137 56. DNase I of tailless nucleosomes and HMGB1-remodeled tailless nucleosomes (4E0). ………………………………………………………… 138 57. Comparison of additional DNase sensitive cuts on the 3 DNA constructs… 140 58. Ava I digestion of 4E0 tailless nucleosomes and remodeled tailless nucleosomes………………………………………………………… 143 xiv

59. Ava I digestion of 2E2 nucleosomes (remodeled tailed and tailless nucleosomes)……………………………………………………… 145 60. Effect of temperature on tailless nucleosomes and remodeled tailless nucleosomes (4E0)…………………………………………………… …… 147 61. Effect of increasing NaCl concentration on mobility of remodeled tailless nucleosomes (4E0)…………………………………….. 149 62. Effect of increasing the levels of unlabeled 161 bp DNA on HMGB1 modified tailless nucleosome………………………………….. … 151 63. Exo III digestion on nucleosomes, HMGB1-remodeled nucleosomes, tailless nucleosome and HMGB1-remodeled tailless nucleosomes………… 153 64. DNase I of tailless nucleosome (4E0) in the absence and presence of 400 nM HMGB1………………………………………………………… 155 xv

Table of tables

1. Comparison of binding of transcription factor to nucleosomes and DNA…… 10 2. The characterization of estrogen receptor α…………………………………. 30 3. Incubation of nucleosomes with increasing HMGB1 concentration………… 43 4. ER binding reactions to nucleosomes in the presence of 400 nM HMGB1…. 45 5. Final concentration of each component of the reaction buffer ………… …… 56 6. Positions of additional DNase I cuts in DNA N’/N”………………………… 99 7. The size of radiolabeled DNA after Ava I digestion and the center of cERE from the labeled end of DNA………………………………………… 104 8. Location of additional DNA bands in N’/N” and N* from DNase I gel...... 139 9. Percent digestion of 4E0 nucleosomes with Ava I………………………… 144 10. Activities for HMGB1 ……………………………………………………. 181 11. Activities for Nhp6 ……………………………………………………….. 182 12. Activities for yFACT……………………………………………………… 183

Appendix

1. Binding of ER to nucleosome in the presence of 400 nM HMGB1 is ATP- independent………………………………………………………………… 194 2. ER binding to 3E1 in the presence of 400 nM HMGB1…………………… 196 xvi

3. ABBREVIATIONS

ACF ATP-utilizing chromatin assembly and remodeling factor

AR Androgen receptor

ATP Adenosine triphoshate

BB Binding buffer

BSA Bovine serum albumin

CTE C-Terminal extension

DB Dilution buffer

DBD DNA binding domain

DTT Dithiotheritol

ERE Estrogen response element

ERα Estrogen receptor-alpha

GR Glucocorticoid receptor

GRE Glucocorticoid response element

HMGB1 High mobility group-1 protein

HRE Hormone response element

KD Dissociation constant

LBD Ligand binding domain

NF-1 Nuclear factor 1 nsDNA Non specific DNA

NPS Nucleosome positioning sequences

N’/N” HMGB1-remodeled nucleosomes

N* HMGB1-remodeled tailless nucleosomes xvii

P-Box Functional region in nuclear receptor that interacts with base pair in DNA

PR Progesterone receptor

RSB Reticulocyte standard buffer

RSC Chromatin structure remodeling

SWI/SNF SWItch/Sucrose NonFermenting

ISWI Imitation SWI

TBE Tris borate EDTA buffer

TBP TATA binding protein

TCA Trichloro acetic acid

TEMED "N,N,N',N'-Tetramethylethylenediamine"

TEN Tris EDTA sodium chloride buffer

TRF1 Human telomeric protein

Tris-HCl Tris hydroxy methyl amino methane hydrochloric acid 1

CHAPTER 1: INTRODUCTION

1. Nucleosome structure and histone/DNA interactions

DNA in eukaryotic cells is packaged in a DNA/protein complex called chromatin.

The basic unit of chromatin is the nucleosome, which is made of 147 bp DNA wrapped

around an octamer of histone proteins in 1.65 turns of a left-handed superhelix (Battistini

et al, 2010; Luger & Richmond, 1998; Segal et al, 2006). Histone proteins are some of

the most conserved proteins known, and there are four histone proteins involved in the

core particle; H2A (13.9 kD), H2B (13.7 kD), H3 (15 kD) and H4 (11 kD). The

nucleosome is disc-shaped, about 11 nm in diameter and 5.7 nm in height and has a

molecular weight of 210 kDa (Lugar et al, 1997). The histone octamer is composed of a

central (H3-H4)2 tetramer flanked on each side by two H2A/H2B dimers and the DNA resides on the outside assembled to form the left-handed superhelix (Luger, 1997;

Wrange, 1995) (figure 1).

There are 3-5 α helices in each of the 4 core histone proteins; but only three take

part in the histone fold; one large and two small with a nonhelical loop region separating

them (figure 2) (Luger, 1997; Zheng and Hayes, 2003). These 3 helices fold into a

secondary structure called the “histone fold”, that is common in all the core histone. The

“histone fold” of H3 and H4 or H2A and H2B interlock to give what is called the

“histone handshake” (figure 3). The “histone handshake” is responsible for the stability in

H3/H4 or H2A/H2B dimers (Luger and Richmond, 1998). The histone proteins interact

with DNA at fourteen different points with the minor groove of DNA through L1& L2

loops and α1 & α2 DNA-binding motifs in between the α helical regions of each of the

core histones (Luger and Richmond, 1998). 2

A B

Figure 1. Nucleosome core particle: ribbon traces for 146 bp DNA phosphodiester

backbones (brown and turquoise) and eight histone protein main chains (blue: H3; green:

H4; yellow: H2A; red: H2B) (A). The views are down the DNA superhelix axis for (A)

and perpendicular to it for the right particle (B). For both particles, the pseudo-two fold axis is aligned vertically with the DNA centre at the top (Luger et al, 1997).

The tetramer of two dimers of (H3/H4)2 binds the central 60 bp of nucleosomal DNA.

The H2A/H2B dimer which is tethered to each half of the histone (H3/H4)2 tetramer organizes 30 bp towards either end of the DNA (Luger et al, 1997). The last 13 bp of

DNA at the entry and exit point of the nucleosome is organized exclusive by the α-helical histone fold extension of H3 and the preceding H3 N-terminal tail (Luger and Richmond,

1998).

The DNA that resides between nucleosome is variable in length and is called linker DNA. The extended structure of polynucleosome array resembles “beads on a string” (Ehrenhorfer-Murray, 2004). Several nucleosomes (about six nucleosomes) are further compacted to form a higher order chromatin fiber of about 30 nm in diameter which is stabilized by the linker histone H1 (Woodcock & Dimitrov, 2001; Luger et al,

1997).

Figure 2. A) Schematic diagram of core histones. Helical regions within nucleosome core are indicated as cylinders and the three helices comprising the histone fold domain in each protein are indicated by the gray box. The red T’s indicate the peptide bond closest to the histone fold domain susceptible to trypsin proteolysis in the nucleosome core. The vertical blue arrows indicate the approximate point where the tails exit either through (H2B and H3) or over/under (H2A/H4) the DNA superhelical gyres to the exterior of the nucleosome (Zheng and Hayes, 2003). B) The histone fold, showing the helix-strand-helix motif (HLH) at each end (Ramakrishna, 1996). 5

Figure 3. Histone handshake motif of H2A/H2B and H3/H4. L1, L2 loops and α1, α2

DNA-binding motif sites are the actual sites on the histone proteins that make contacts with the DNA. A) The histone-DNA contacts of the H3-H4 tetramer. H3-H4 binds the central 60 bp of nucleosomal DNA. B) The histone-DNA contacts of the H2A/H2B dimers. H2A/H2B binds 30 bp of nucleosomal DNA (Lugar et al, 1997).

1.1 Histone tail domains

The core histone proteins are composed of a high percentage of positively charged amino acids, arginine and lysines (figure 2).The core histone tail domains are located at the N-terminal of each of the core histones; in addition to the C-terminal of histone H2A

(Hayes et al, 1991) as shown in figure 2a. Each of the four core histones of the nucleosome has about 15-to 45 positively charged amino acids in the N-terminal tail domain (Protacio et al, 2000; Wolffe &Hayes, 1999).

Specific amino acid residues in the N-terminal and C-terminal domains of histones (lysine, histidine, serine, arginine, threonine) are known to undergo posttranslational covalent modification such as phosphorylation, methylation and acetylation (Li et al, 2002). Such modifications are thought to play an important role in chromatin function and gene transcription (Wong et al, 2002). The combinatorial nature of the modifications that take place in histone tail domains is termed the “histone code hypothesis” (Jenuwein et al, 2001). For example, the acetylation of multiple lysine residues of histone H3 and H4 by enzymes known as histone acetyltransferases (HATs) is required for DNA replication (Unnikrishna et al, 2010). On the other hand, the cleavage of the acetyl groups by enzymes known as histone deacetylases (HDACs) is associated with transcriptionally inactive chromatin. Therefore, the histone tail domain plays a big role in epigenetic regulation.

1.2 Tailless histones

The exposed tails can be cleaved off with trypsin without affecting the stability of the nucleosome (Yang et al, 2005). Figure 2 illustrates where the trypsin cuts occur. In the x-ray crystal structure, the tail domains are unstructured and not observed. The tails extend or exit the nucleosome through minor groove of the DNA to the exterior. The tail domains of H2A and H4 pass through the minor grooves from above and below the top and bottom of superhelical turns whereas the tail domains of H3 and H2B passes through the minor grooves amid the adjoining DNA superhelical gyres (White et al, 2001; Luger

et al, 1997). The C-terminal tail domains of H2A exit the nucleosome at the dyad (White

et al, 2001). Tailless nucleosomes have been used as a model for highly acetylated

nucleosomes by some labs (Polach et al, 2000). The reason being that, acetylation of each

of the basic amino acids reduces the positive charge by one unit. This might weaken any

interaction between the highly negatively charged DNA and the positively charged histone tail domains. Histone acetylation has been suggested to be an important initial

step in the activation of a gene, required to facilitate subsequent access of transcription

factors to sites in nucleosomal DNA (Li et al, 2002; Wong et al, 2002; Unnikrishna et al,

2010). Hyperacetylation therefore, reduces the positive charge on the histone tail

domains, which is comparable to cutting the tail domains off. The rate of transcription

elongation through a nucleosome was comparable in nucleosomes without tail domains

and those that had the tail domains acetylated (Protacio et al, 2000).

2. Chromatin and remodeling 8

For transcription to take place in chromatin, RNA polymerase II, transcriptional activators, and general transcription factors must interact with their binding sites in the regulatory region of DNA. In all eukaryotes, however, the genome is packaged in nucleosomes, which restricts binding of the general transcription factors and therefore gene expression (table 1) (Ding et al, 1997; Galati et al, 2006; Chen et al, 1994;

Imbalzano et al, 1994 and 1998). For example, the binding of TATA –binding protein

(TBP) to the TATA sequence is severely inhibited (by a factor of 105) when the DNA is incorporated into a nucleosome (Imbalzano et al, 1994). Also Nuclear Factor 1 (NF-1) binding to its recognition sequence in a nucleosome is reduced by 100-300-fold compared to free DNA (Blomquist et al, 1999).

Our laboratory was interested in using one DNA construct with the different hormone response elements for estrogen receptor (ER), progesterone receptor (PR) and glucocorticoid (GR) in order to make a direct comparison of these various receptors and how they bind to free DNA, and DNA within a nucleosome. We have previously shown that ER binding to nucleosomes is reduced by a factor of 50 compared to binding in naked DNA (Sarpong, 2006). Progesterone receptor (PR), on the other hand, did not bind to nucleosomes even at 150 nM of PR under our conditions. In contrast, it was reported that PR binds to a nucleosome (Pharm et al, 1992). Glucocorticoid receptor was a special case, because it bound to almost all the DNA constructs, including those that did not have a glucocorticoid response element (GRE) (Sarpong, 2006, appendix). A comparison of transcription factors binding to nucleosome is presented in table 1. 9

Figure 4. Nucleosome core particle. A) Histone tails between DNA gyres. The H2B

(red) and H3 (blue) N-terminal tails pass through channels in the DNA superhelix (white) formed by aligned minor grooves. H4 is green and H2A is yellow. B) ribbon traces for

146 bp DNA phosphodiester backbones (grey) and eight histone protein main chains

(blue: H3; green: H4; brown: H2A; red: H2B). The histone tail domains extend from the nucleosome core. (http://lugerlab.bmb.colostate.edu/Gallery.html) 10

Table 1. Comparison of binding of transcription factor to nucleosomes and DNA

Transcription Relative binding affinity to References factors nucleosomes GR Binds to nucleosomes with similar Pina et al., 1990; Li and binding affinity as in DNA. Wrange, 1995; Perlmann, 1992 Reduced 2 fold ERα 100 fold decrease upon nucleosome Ruh et al., 2004, Sarpong, 2006 binding NF-kB Binds to nucleosome with same Angelov et al., 2004 affinity as to DNA TRF1 10-20 fold decrease upon Galati et. al., 2006 nucleosome binding Sp1 10-20 fold decrease upon Li et al., 1994 nucleosome binding PR Strong binding Pharm et al., 1992 PR Little or no binding upon Sarpong, 2006 nucleosomes GAL4 10-100 fold decreasing to Chen et al., 1994; Taylor et al., nucleosome binding 1991; Burns et al, 1997 TBP (105) decrease in nucleosome Imbalzano et al., 1994 binding NF-1 100-300 fold decrease upon Blomquist et al., 1999 nucleosome TFIIIA Decreased upon nucleosome Vitolo et at., 2004; Lee et al, binding 1993

Abbreviations are: ERα, Human estrogen receptor-alpha; GR, Glucocorticoid receptor; PR, Progesterone receptor; TBP, TATA binding protein; TRF1, telomeric protein; NF-1, Nuclear factor 1; GAL4, galactose induced transcription factor. 11

2.1 ATP-dependent chromating remodeling

Nature has devised several ways for transcription factors to overcome the barrier

imposed by incorporation of DNA into nucleosomes. A mechanism recognized to

remodel nucleosomes utilizes an ATP-dependent mechanism in which DNA can be

translocated relative to the histone proteins by chromatin remodeling complexes

(Phelan et al, 2000). These complexes contain an essential motor protein that catalyzes

ATP hydrolysis to provide energy for translocation. The SWI/SNF (SWItch/Sucrose

NonFermentable) complex is a 2 MDa chromatin remodeling complex composed of 11

different polypeptide subunits. The catalytic subunit is SWI2/SNF2, which contains the

ATPase activity. The homolog in humans is hBRM (hSNF2α) or BRG1 (hSNF2β) (Hill

and Imbalzano, 2000). SWI/SNF has been shown to alter the rotational phasing of DNA

in nucleosome and enhance binding of TBP to TATA site (Imbalzano et al, 1994),

increase binding of transcription factor (GAL4) to nucleosomes (see table 1, Burns et

al, 1997) and increase restriction enzyme (Xho I & Rsa I) access to their recognition

sites buried in a nucleosome (Kassabov et al, 2003; Schnitzler et al, 1998). Chromatin

structure remodeling (RSC) complex, a SWI/SNF related chromatin remodeling

complex from yeast, also alters nucleosome structure in an ATP-dependent manner

(Lorch et al, 1998).

2.2 ATP-independent chromatin remodeling

There is one complex in yeast that alters nucleosome structure and does not require ATP to remodel nucleosome structure. These proteins appear to have no enzymatic activity and do not translocate DNA. In Saccharomyces cerevisiae, this 12

heterotrimeric complex is called yFACT. The complex consist of three proteins; Nhp6,

Pob3 and Spt16 (Formosa et al, 2001). The Nhp6 unit contains a single DNA binding motif similar to those found in members of the HMG box family of DNA binding proteins (Ruone et al, 2003). yFACT has been shown to enhance the sensitivity of specific sites within nucleosomal DNA to DNase I (Formosa et al, 2001). Nhp6 proteins, can independently convert a canonical nucleosome into another form that is more accessible to DNase I cleavage at specific sites (Rhoades et al, 2004). Human FACT

(Spt16/p140 and SSRP1) stimulates transcriptional elongation by RNA pol II through histone H2A/H2B dimer displacement in the nucleosome while suggesting that the hFACT remodels nucleosomes as shown by yFACT ( Mason and Struhl, 2003).

3. HMGB1

High mobility group 1 (HMGB1) is a highly conserved protein that is found in all mammalian cells (Bustin et al, 1990). The protein has a molecular weight~25kDa, and is made up of 215 amino acid residues. Figure 5 shows the schematic representation of

HMGB1 protein. The protein can be divided into three separate domains; A-box (1-89),

B-box (91-176) and C-terminal domain (185-215). The A- and B-domains are highly positively charged due to a high content of basic amino acid residues (arginines and lysines) (Bustin et al, 1990). These domains are known to interact non-specifically with

DNA (Bustin et al, 1990). The C-terminal contains 30 residues, which are all negatively

charged acidic amino acid residues of aspartic and glutamic acids. This part of the

HMGB1 is absolutely essential for the formation of a stable complex between the TBP

and HMGB1 when bound to the TATA box in DNA (HMGB1/TBP/TATA complex) 13

(Das and Scovell, 2001). The C-terminal of HMGB1 also interacts with histone H3 in a

nucleosome and competes with H1 at the linker region between nucleosomes (Kawase et

al, 2008; Varga-Weisz et al, 1994).

Figure 5. Domain structure of HMGB1. HMGB1 contains 3 domains; the A and B

boxes are basic in nature and contains a large percentage of arginine and lysines; and the

C-terminal is acidic in nature and contains only aspartic and glutamic acid (Das and

Scovell, 2001).

3.1 Effect of HMGB1 on transcription factor binding

HMGB1 binds nonspecifically to the minor groove of DNA and bends the DNA

or makes the DNA more flexible (Zhang et al, 2009; Nardulli et al, 1998; Agresti &

Bianchi, 2003). It is this flexure function of HMGB1 that we will propose aids in the binding of transcription factor to nucleosomes. HMGB1 enhances RNA pol II

transcription in a subset of genes, including those regulated by steroid hormone receptors.

HMGB1 enhances binding of PR and ER to DNA (Onate et al, 1994; Das et al, 2004).

Coexpression of HMGB1 or HMGB2 increased PR-mediated transcription in mammalian 14

cells by as much as 7- to10-fold without altering the basal promoter activity of target

reporter genes (Onate et al, 1994).

HMG-box containing proteins, such as yeast BAF57 in SWI/SNF, have been

shown to enhance binding of transcription factors to the nucleosomes (Guyon et al,

1999). BAF57 subunit in the SWI/SNF complex has been reported to regulate ER

function in breast cancer cells (Garcia-Pedrero et al, 2006) and is required in mice for

nucleosome remodeling in vivo (Chi et al., 2002). In yeast, the HMGB1- related Nhp6 proteins facilitate the interaction of the heterodimeric Spt16/Pob3 proteins with nucleosomes (Brewster et al, 2001; Formosa et al, 2001).

ATP-utilizing chromatin assembly and remodeling factor (ACF) is an ATP-

dependent nucleosome remodeling complex with imitation SWI (ISWI) as the ATPase

subunit. This complex can remodel nucleosomes by sliding of histone octamers on DNA

through the hydrolysis of ATP. HMGB1 accelerates the sliding activity of ACF when it interacts with nucleosomal linker DNA overlapping ISWI-binding sites (Bonaldi et al,

2002). HMGB1 has been shown to enhance the activity of chromatin remodeling

complexes, SWI/SNF and RSC, by enhancing the kinetics of sliding of histone octamers

relative to the DNA by ~2-fold (Ugrinova et al, 2009).

The acidic tail of HMGB1 has been shown to interact specifically with Lys36 and

Lys37 in histone H3. These interactions are important for modulation of the DNA and

chromatin binding activities of HMGB1, as well as biological functions of HMGB1

(Kawase et al, 2008).

3.2 Effect of HMGB1 in DNA repair and V(D)J recombination 15

HMGB1 has also been shown to be an important protein in DNA repair pathways.

HMGB1 binds to cisplatin lesions and inhibits nucleotide excision repair in vitro (Huang et al 1994). V(D)J recombination is a pathway that is initiated by deliberate double strand breaks in DNA at specific areas in the genome in B and T cells. This pathway is important in creating immunoglobulin diversity in B and T cell receptors (Swanson,

2004). V(D)J recombination is regulated by the activity of RAG1 and 2 proteins, cleaving the DNA at specific recombination signal sequences. HMGB facilitates the assembly of

RAG1/2 recombination signal sequences (RSS) DNA complexes that are components for

V(D)J cleavage (Lange and Vasquez, 2009; Dai et al, 2005) and has also been shown to stimulate cleavage and RAG protein binding to DNA (Gent et al. 1997). The DNA binding domains in HMGB1 were found to be essential in assembling RAG1/2 RSS complex and stimulating DNA cleavage in nucleosomes (Dai et al, 2005).

Altogether, these reports show that the HMG box proteins can play a major role in chromatin remodeling in increasing the binding affinity of transcription factors, repair enzymes and the recombination machinery within chromatin.

3.3 Effect of HMGB1 in transcription

HMGB1 has also been shown in numerous studies to stimulate transcription by increasing the sequence-specific binding affinity of steroid hormone receptors to its hormone response elements (Boonyaratanakornkit et al, 1998; Melvin and Edwards,

1999; Zhang et al, 1999). For example, it facilitated PR mediated transcriptional activity

(Onate et al, 1994; Prendergast et al, 1994). HMGB1 protein stimulates transcription by

RNA polymerase II and III (Tremethick and Molloy, 1988). Joshi also showed that transfected HMGB1 stimulates gene expression of ER mediated transcription in ER- 16

negative human osteosarcoma cells (Joshi, 2010) that also had ER expression vector transfected. HMGB1 deficient mice show a distinct phenotype and die shortly after birth

due to hypoglycemia and were deficient in the activation of GR responsive genes

(Calogero et al, 1999), which suggests that HMGB1 is important in transcriptional

regulation of specific genes.

HMGB1 was reported to up-regulate the human topoisomerase IIα gene (Stros et al, 2009). During transcriptional activation of pS2 gene, H1 is replaced by HMGB1/2 in a nucleosome that has the ER binding site. This is important because HMGB1/2 may be acting as activator and may also stabilize the binding of ERα to binding sites on DNA (Ju et al, 2006).

4. Estrogen receptor (ER)

ER belongs to the superfamily of nuclear hormone receptors that regulate gene expression by binding to specific palindromic sequences called hormone response elements (HREs) (Melvin et al, 2004). Steroid hormone receptors are ligand-dependent and bind as homodimers to response elements in the enhancer regions of specific genes

(Onate et al, 1994). Two subtypes of ER are expressed in mammalian cells; ERα and

ERβ. In this study ERα is referred to ER.

The estrogen receptor has five structural/functional domains: 1) The A and B domains (N-terminal domain) are required for maximum transcriptional activity, and the domain controls transcription in a cell-specific manner (Melvin et al, 2004). 2) The C domain is the most highly conserved and also, the DNA binding domains (DBD).

Crystallography studies have shown that ERα DBD binds to EREs as a homodimer 17

(Schwabe et al, 1993).The DBD consists of a highly conserved core with two distinct zinc fingers and a 30 amino acid segment, termed the C-terminal extension (CTE).

Within the core DBD, α-helix 1 extends between the two zinc fingers and makes base

specific contacts in the major groove of the HRE DNA. The second α -helix (helix 2) does not contact DNA but is important for the overall folding of the core DBD. 3) D, hinge regions. 4) E domain is a ligand binding (LBD) region that is conserved throughout evolution and contains activation function-2 (AF-2). 5) F domain is responsible for distinguishing between estrogen agonists versus antagonists, through interaction with

cell-specific factors (Montano et al, 1995) (figure 6 & 7).

Intracellular estrogen receptor is activated in the presence of estrogen (E2). The

activation process involves a change in conformation of the ER protein, which involves

the dissociation of hsp70, hsp90, and other proteins from monomeric ER in the nucleus

(Klinge et al, 1997). ER then dimerizes and binds to a recognition sequence

(AGGTCAnnnACTGGA) as a homodimer to stimulate transcription (Devin-Leclerc et al,

1998; Klinge, 2001). ER has also been shown to interact with other transcription factors bound to DNA and stabilizes that DNA/transcription factor interaction and recruits other transcription factors to the site (Klinge, 2001). ER tethers to Sp1 in conferring estrogen responsiveness on regulatory sequences in the uteroglobin gene (Scholz et al, 1998), and to blc-2 gene in breast cancer cells (Dong et al, 1999).

5. Nucleosome positioning sequences, translation positioned and rotationally phased

nucleosomes

An artificial nucleosome positioning sequence (NPS) is the repeating motif

containing [5’-(A/T)3 NN(G/C)3 NN-3’]. This sequence has been shown to have a high

affinity for binding the histone octamer and bending it to accommodate a strong DNA-

histone interaction in a nucleosome (Shrader & Crothers, 1989). This therefore will fix

the DNA on the histone octamer in a rotationally phased and translationally positioned manner.

We used NPS to create a homogenous population of nucleosomes that were translation positioned and rotationally phased. The translational position is the position of the ends of the DNA sequence relative to the nucleosome, while the rotational phase has to do with the orientation of the major and minor grooves relative to the surface of histone octamer (Woodcock and Dimitrov, 2001). 19

Figure 6. Schematic representation of the functional domain organization of nuclear receptors. A/B, transactivation domain (AF1); C, DNA binding domain

(DBD); D, hinge regions; E, ligand binding domain (LBD); F, important domain for

ERα function. The enlarged part of the DBD shows the core DBD and the C-terminal extension for the human estrogen receptor alpha (hERα) (Ruff et al, 2000; Melvin et al, 2004). 20

Figure 7. ERα DBD binding to cERE. The picture is a crystal structure showing the helix 1 of ERα DBD interacting with the major groove of DNA. Two ER DBDs interact to form a dimer at AGGTCAnnnACTGGA.

Objectives

It has been reported that ER binds to cERE in a nucleosome in a dose-dependent manner, and that HMGB-2 increases the binding affinity of ER to a nucleosome (Ruh et al, 2004). However, the gel electrophoresis data do not appear to agree with their stated interpretation. Also nuclear extract was used as the source of ER, which may contain other proteins contributing to both specific and non-specific binding ER to nucleosomal

DNA. The nucleosome fractionation from sucrose gradient centrifugation is also contaminated with free DNA. Therefore, it becomes difficult to determine if the binding observed is due to ER binding to free DNA or to the nucleosomal DNA (Ruh et al, 2004; figures 1a and 1b). Our laboratory uses purified ER in the binding studies. We were also able to isolate pure rotationally phased and translationally positioned nucleosome fractions which had no free DNA cofractionating with the nucleosomes. Therefore, we are able to quantify the characteristics of pure phased nucleosome fractions and for ER binding.

The main objectives of this dissertation are to determine:

1. The influence of HMGB1 on remodeling nucleosome structure

2. The effect of HMGB1 on ER binding to remodeled nucleosome

3. The effect of removing the histone tail domain on remodeled nucleosomes and the

effect of HMGB1 on access to enzyme and ER

CHAPTER 2: MATERIALS AND METHODS

1. Construction of plasmids containing cERE

Water used in all solution preparations was Millipore water from Barnstead E-pure,

PSLB 520. All DNA molecules used in the experiments were prepared by Dr Ron

Peterson, Ohio Northern University). Artificial nucleosome positioning sequences have

been shown to bend easily when constructed such that segments consisting of A&T or

G&C are separated by 2 base pairs. Multiple repeats of the (A/T)3NN(G/C)3NN motif

were found to be more effective than natural sequences at positioning nucleosomes

(Shrader and Crothers, 1989).

To create a set of DNAs in which a DNA sequence, such as cERE, could be inserted

at different positions in the DNA, along with the nucleosome positioning sequence in a

single step, and to ensure that the insert is placed in a particular direction, we made use of

a restriction enzyme (RE) that has an asymmetric cutting site with some redundancy. The

enzyme, Ava I, recognizes and cuts within the C/PyCGPuG where / represents the cutting

site (Py is a pyrimidine, and Pu is a purine). Sequences that are cut by Ava I include

CCCGGG, CTCGGG, CCCGAG & CTCGAG. The DNAs were constructed by replacing

the Ava I site in pGEM3Zf(+) with the sequence CTCGGG and cloning appropriate

oligonucleotide between two of the other restriction sites.

Figure 8 A illustrates how Ava I cuts the plasmid DNA used to construct the 161 bp

DNA. The plasmid is first cut with Ava I and a nucleosome positioning sequence (NPS)

inserted. The inserts are in two orientations. Only one of the orientations can ligate

perfectly with the plasmid DNA. After the first cut with Ava I, the orientation 1 fragment

is the only product that can be formed. The ‘N’ at the end of orientation 1 fragment 23

stands for either a GC or TA bp. If there is a GC to the left of the fragment, a new Ava I is generated after inserting the fragment into the plasmid. An AT bp at the end will not generate an Ava I site. The orientation 2 is not able to generate an Ava I site, and therefore, can not be ligated to another DNA cut with Ava I. This ensures that the insert is placed in only one direction.

The four nucleosome positioning sequences in each of the DNA constructs were built up by sequentially cloning fragments A or B (fig 8B) in one orientation so that an Ava I site was regenerated at only one side of the insert fragment. A set of plasmids with unique Ava I site at different translational positions within the nucleosome positioning sequence are generated by sequential cloning of the nucleosome positioning sequences.

Figure 8 C illustrates how a single Ava I site is generated each time the nucleosome positioning sequence is inserted into a construct. This allows for the positioning of ERE at different translational positions within the context of the nucleosome positioning sequences (modified from Shrader & Crothers 1989; Li & Wrange 1993; Li & Wrange

1995). The figures below illustrate how the plasmids with cERE at different translational positions were constructed.

Plasmid DNA CTCGGG A GAGCCC

Cut with Ava I

C + TCGGG

GAGCC C

Ligation CCGAN N TCGGN N N NGGCT N NAGCC Insert fragment Insert fragment orientation 2 orientation 1 No product CTCGGN NTCGGG GAGCCN NAGCCC

Frag ment can insert in only one orientation

Figure 8 A. Ava I restriction site in plasmid. The scheme shows the Ava I restriction sites and the insertion of nucleosome positioning sequences. The “N” represents either

G/C or T/A bp as shown in fig 8 B. If N/N = T/A then the site is not Ava I site. If N/N =

G/C then the site is Ava I site.

Figure 8 B. Nucleosome positioning sequences. Fragments A and B are DNA fragments for insert of a nucleosome positioning sequence with a single Ava I site. The overhanging ends can ligate directionally to an asymmetric Ava I site on a plasmid DNA.

The ends marked with (+) will generate an asymmetric Ava I site when it is joined to 25

another (+) end. The (-) ends will not regenerate an Ava I site when joined to either type

of ends.

Figure 8 C. Construction of sets of plasmids with unique Ava I site. This procedure produces three plasmids with an asymmetric Ava I site located at the end, in the middle or

20 bp from one end of the nucleosome positioning sequences that is 80 bp long. 26

1.1 Construction of DNA 4E0

The plasmid was first cut with Ava I to generate TCGG at one 5’ end of one strand and

AGCC at the 5’ end of the other strand. Because the Ava I has an asymmetric cutting site, the plasmid will ligate to a piece of DNA cut with Ava I in only one direction. Fragment

A contains TCGG which can base pair and ligate AGCC end of the plasmid. The other end of the insert generates an Ava I site. The Ava I site is cut again and another fragment

A inserted. A total of four fragments A are inserted into the plasmid. Finally, a cERE which had been synthesized to contain TCGG and AGCC was inserted into the plasmid.

This DNA construct therefore, contains four nucleosome positioning sequences next to a cERE. This construction places the ERE 40 bp from the dyad axis.

1.2 Construction of DNA 2E2

The plasmid was first cut with Ava I to generate TCGG at one 5’ end of one strand and

AGCC at the 5’ end of the other strand. Two of fragment A was sequentially cloned into the construct as before. Two of fragment B was sequentially cloned into the construct as before. This creates an Ava I in middle of the 4 nucleosome positioning sequences. A cERE which had been synthesized to contain TCGG and AGCC was inserted into the plasmid. This DNA construct therefore, contains two nucleosome positioning sequences on each side of the cERE.

2. Isolation and purification of HMGB1 & HMGB2 proteins

2.1 Isolation of calf thymus nuclei

The proteins were isolated under non denaturing conditions using differential

precipitation with ammonium sulfate and HPLC using a Pharmacia Mono Q HR 5/5

anion exchange column. A 200 g of calf thymus (Bellville Market, Bowling Green, OH)

was homogenized in buffer A (50 mM NaCl, 20 mM Tris-HCl (pH 7.6), 1 mM EDTA,

10% (w/v) glycerol, 5 mM BME and 0.5 mM PMSF). The thick homogenate was filtered

through two layers of cheesecloth. The filtrate was then transferred into 50 mL

centrifuge tubes, and centrifuged at 14000 x g for 40 mins. The supernatant were

discarded and the pellet washed repeatedly with buffer A until no observable lipid

remained.

2.2 Purification of HMGB proteins

The pellet was resuspended in two volumes of buffer B (20 mM Tris-HCl (pH

7.6), 5 mM BME and 0.5 mM PMSF). A 0.1 volume of 1.65 M ammonium sulfate (AS)

was slowly added, while stirring with a magnetic stirrer. The AS solution was prepared

with buffer B. The suspension was stirred gently for 30 mins at 4 oC and then sedimented

at 13000 x g for 20 mins. The final concentration of the supernatant was made to 2.6 M

AS by the slow addition of solid salt. The solution was stirred gently for an hour at 4oC and sedimented at 100,000xg for 20 mins, using the Beckman ultracentrifuge, SW28 rotor. The clear supernatant was transferred into ¾ inch dialysis tubing (10kDa MWCO).

The AS was dialyzed from a high concentration of 2.6 M to µM concentrations in multiple (6) transfers into 4L of buffer (20 mM Tris-HCl (pH 7.6), 5 mM BME and 0.5 mM PMSF). Dialyzed solution that contained HMGB proteins was concentrated to 10 28

mL by ultrafiltration (10 kDa MWCO, Diaflow by Amicon) using N2 pressure unit at 60

psi. The homogenate was further concentrated to 5 mL, using Millipore centrifuge concentrator tubes (10 kDa MWCO, 1600 rpm, 219 rotor at 4 oC).

An aliquot of 5 M NaCl was added to the concentrated protein solution to a final concentration of 0.8M NaCl. This precipitates out extraneous proteins with might precipitate and block the column during the HPLC. The solution was gently stirred for an

hour and sedimented at 12,000 xg for 20 mins. The supernatant was transferred into

another dialysis tube (10kDa MWCO) and dialyzed against buffer B. The solution was concentrated to one mL using Millipore centrifuge concentrator (10kDa MWCO). The concentrated protein solution was filtered using sterile Acrodisc filter (pore size 0.2 microns). The two HMGB proteins (HMGB1 and HMGB2) were separated from other proteins and each other by a linear NaCl gradient. The high salt buffer contained 0.8 mM

NaCl, 20 mM Tris-HCl (pH 7.6) and 5 mM BME. The low salt contained 0.2 mM NaCl,

20 mM Tris-HCl (pH 7.6) and 5 mM BME). The computer program was set up to run the gradient containing a 2 minute wash with low salt buffer, to remove unbound proteins; an

18 minute linear salt gradient was run to elute and fractionate HMGB1 and HMGB2.

After HMGB1/2 elution, a 2 minute wash with high salt buffer and finally a 4 minute wash with low salt buffer were done. The HMGB2 & HMGB1 were eluded at 13.03 and

14.05 minutes respectively, on a linear salt gradient.

Following the collection of samples, the HPLC unit and the column was washed with distilled water for 4 hrs at 1 mL/minute and then at 0.1 mL per minute overnight.

This was required to remove residual salts since residual salts in the HPLC will corrode

and damage the tubing. The Mono Q column was stored in 20% ethanol by running the 29

solvent for about 10 minutes, then the system was shut off and the column stored by

covering both the ends properly with suitable caps. This work was performed in Dr. D.

Diagram’s lab (University of Toledo, Health Science Campus) with his assistance, which we gratefully acknowledge.

2.3 Characterization and Storage of HMGB1 and HMGB2 Proteins

The HMGB1 and HMGB2 fractions were dialyzed into HMGB1 working buffer

(160 mM NaCl, 10 mM Na2HPO4, 1 mM DTT at pH 7.3) and stored in 50 μL aliquots at

-80 oC. The fractions were analyzed on SDS-PAGE to determine the purity of the

proteins. A single band corresponding to ca. 25 kDa molecular weight was observed for

both HMGB1 and HMGB2. The concentration was determined using an extinction

coefficient of 20,500 M-1 cm-1 at 280 nM (Chow et al, 1995). The amount of HMGB1

and HMGB2 recovered were usually about 3.0 mg and 2.8 mg, respectively. The

concentration of HMGB1 and HMGB2 were adjusted to ~ 1 µg/µL in HMGB working

buffer.

3. Estrogen receptor alpha (ERα)

The recombinant full length human ERα is 66.4 kDa, 559 amino acid residues and

was purchased from Invitrogen (Cat #, P2187). The protein was expressed and purified

from recombinant baculovirus-infected insect cells. The purity was greater than 80% as

determined by a Coomassie blue-stained SDS-PAGE mini gel by Invitrogen. The ERα

was stored in storage buffer containing 50 mM Tris-HCl (pH 8.0), 500 mM KCl, 2 mM

DTT, 1 mM EDTA, 1 mM sodium orthovanadate, 10% glycerol. 30

Table 2. The characterization of estrogen receptor α.

Concentration Specific activity Functional receptor Protein Lot No. (mg/ml) (pmol/mg) concentration (nM)

ERα 26467A 0.26 6923 2088

The table shows the lot number, total protein concentration, specific activity and functional receptor concentration for each of the lots used in this research.

4. Antibodies to core histone proteins

The anti-histone antibodies (H2B, H3, and H4) were purchased from Active Motif.

Histone H2B (Cat # 39125) has a molecular weight of 15 kDa with the epitope being the peptide including the C-terminal region of histone H2B. Histone H3 (Cat # 39163), molecular weight 17 kDa, with the epitope being the C-terminal peptide of histone H3.

Histone H4 (Cat # 39269) molecular weight 8 kDa with the epitope being the human histone H4. Histone H2A (Cat # 07-146) was purchased from Millipore, molecular weight was 14 kDa. The epitope is KLH-conjugated synthetic peptide corresponding to amino acids 88-97 (IRNDEELNKL) of human histone H2A. 31

5. Preparation of oligonucleosomes

5.1 Isolation of chicken erythrocytes (CE) nuclei

Chicken blood was purchased from Pel-Freez, Arkansas. About 500 mL of chicken blood in citrate buffer was centrifuged at 650 x g for 5 minutes to separate the

plasma for the cells. The cells were then washed five times with isolation buffer A

(0.85% NaCl, 0.01% methiolate, 6 mM Na2HPO4 pH 7.0) and pelleted at 650 x g for 5

minutes. The supernatant was then discarded. About 250 mL of suspended cells were

recovered at the end of the centrifugation. Aliquots of 10 mL were suspended with 10%

glycerol and stored at -80 0C.

About 15 mL of cell suspension was thawed to 4 oC and homogenized in a glass

homogenizer with Teflon pestle for 30 seconds at 4 oC in 25 mL of reticulocyte standard

buffer (RSB) (10 mM Tris-HCl (pH 7.2), 10 mM NaCl, 3 mM MgCl2 containing 0.5%

IGEPAL, nonionic detergent) to lyse the cell membrane. The solution was made 0.2 mM

PMSF (protease inhibitor) from 0.2 M stock in isopropanol immediately before use. The

resulting suspension was centrifuged at 5000 rpms SS34 rotor for 5 minutes at 4 oC to

pellet the nuclei. The nuclei were washed with RSB buffer containing PMSF and

IGEPAL by suspending in a total volume of 25 mL. The subsequent washes were

without the detergent and contained only buffer/PMSF.

The CE nuclei were washed twice with 20 mL of RSB buffer and centrifuged at

5000 rpms and the supernatant discarded. The cells were finally washed with RSB alone

and the final CE nuclei pellet was resuspended in a total of 5 mL with RSB buffer.

The final concentration in terms of DNA was determined by measuring the A260 of nuclei

dissolved in 2 M NaCl / 5 M urea, with the same solution as the blank. Double stranded 32

DNA concentration was calculated by assuming 1 O.D. at 260 nm was equivalent to 50

µg DNA per mL. The preparation yields 5 mL of CE nuclei suspension and a DNA

concentration of 15,000-16100 µg DNA per mL (300-330 O.D. units).

5.2 Preparation of H1-free oligonucleosomes

H1-free oligonucleosomes from chicken erythrocyte (CE) were used as a source of core histones in the preparation of nucleosomes. CE nuclei were isolated as above except that additional washes were made with a buffer B (15 mM Tris-HCl, (pH 7.5), 15 mM NaCl, 15 mM β-mercaptoethanol, 60 mM KCl, 0.5 mM spermidine. 0.15 mM spermine) containing 0.34 M sucrose. The concentration of nuclei (5 mL total), was

adjusted to 5000 µg DNA / mL and 50 µL of 100 mM CaCl2 was added to a final

concentration of 1 mM. The solution was incubated at 37 oC and then 75 U/mL

micrococcal nuclease (Worthington, Cat # X5C7939) was added, followed by 4 min

incubation at 37 oC. The reaction is stopped by adding 100 mM EDTA, to a final

concentration of 2.2 mM. The nuclear suspension was then centrifuged at 4000 x g for 5

min at 4 oC. The supernatant is discarded and the nuclear pellet is suspended in 5 mL of

0.2 mM EDTA. Then 810 µL of 4 M NaCl was added slowly while vortexing to make a

final concentration of 0.65 M NaCl. The suspension was homogenized, and re-

centrifuged for 5 min at 4000 x g at 4 oC. At this step, the nuclei lyse. The pellet formed

after centrifugation is viscous and is discarded. The supernatant (5 mL) that contains

soluble donor oligonucleosomes was transferred into a fresh tube. It is important to keep

the NaCl concentration at 0.65 M in order to dissociate histone H1, while the core

histones remain bound to the DNA in nucleosome at this NaCl concentration. 33

The supernatant was transferred onto a Sepharose CL-4B (Sigma, Cat #

CL4B200) column (2.5 x 25 cm) equilibrated with a buffer containing 0.65 M NaCl, 5 mM Tris-HCl (pH 7.5), and 0.2 mM β-mercaptoethanol added just before use. The column was eluted with a flow rate of about ten drops per minute with ca. 2.5 mL collected per tube at 4 oC. An SDS gel was run to determine the purity of the proteins in

the fractions. The fractions that contained only the four core histones were pooled

together and dialyzed (10kDa MWCO) at 4 oC overnight in TE buffer. Aliquots were

stored at -80 oC.

5.3 Preparation of tailless oligonucleosomes

The H1-depleted CE oligonucleosomes of (1 mg/mL) was treated with a final

concentration of 0.5 µg/mL trypsin, for 30 min at 37 oC and the reaction stopped by adding a trypsin inhibitor (Sigma, T9003) to a final concentration of 10 µg/mL (Mutskov et al, 1998). The cleaved tail domains, inhibitor and trypsin were separated from the rest of the oligonucleosomes with Sepharose CL-4B column. An 18% SDS gel of the tailless

histones was run along with core histones to test the purity of the histone proteins. The

gel was stained with Coomassie blue. The A260 was measured for the initial fractions. The

initial fractions with high A260 contained the tailless oligonucleosomes. These fractions

were pooled together and dialyzed in TE buffer. Aliquots were stored at -80 oC.

6. Sequence of DNA

The 15 bp synthetic ERE 5’ AGGTCActgTGACCT 3’ (purchased from IDT) was placed at the middle (2E2; at dyad axis) of the 161 bp DNA segment or 40 bp off the dyad axis (4E0) in the plasmid pGEM3Zf(+). A 161 bp 2E2 DNA fragment had the cERE flanked with two NPS at 5’ end and two NPS at 3’ end of the cERE and the 4E0 had the cERE with 4 NPS at 5’ end of the cERE. Nucleosome positioning sequence

(NPS) is a 20 bp repeat of: 5’-TCGGTGTTAGAGCCTGTAAC -3’ (Li and Wrange,

1995).

The complete sequence of the DNA fragment from SP6 promoter primer to T7 promoter primer of a p2E2-pGEM3Zf (+) and p4E0-pGEM3Zf (+) plasmids are shown in the fig 9 and 10, respectively. This fragment contains a 161 bp DNA within the EcoR I and Hind III restriction sites. The15 bp cERE was placed within the DNA such that it is translationally fixed within the major groove facing outward, for optimum binding when the DNA is incorporated into a nucleosome. The orientation of the major groove of the

DNA is established by the constraints imposed by the binding characteristics of the NPSs.

Figure 9. DNA sequence for 2E2. The 2E2 DNA fragment has 2 nucleosome positioning sequences, a repeat of 5’-TCG GTG TTA GAG CCT GTA AC -3’ (Li and

Wrange, 1995) (starting at the 28th bp, counting from the cleavage site of EcoR I) on each side of a cERE, 5’AGGTCActgTGACCT 3’. The pink shaded nucleotides represent the AT-rich region that are constrained so that the minor groove faces toward the histones and the major groove is directed and exposed to the outside. Green shaded nucleotides are the GC-rich regions that have their major groove facing inward toward the histone core. The symbol (•) depicts the center of ERE and (*) depicts the center of the DNA fragment produced by EcoR I and Hind III double digest. 36

Figure 10. DNA sequence for 4E0. The 4E0 DNA fragment has 4 nucleosome positioning sequences, a repeat of 5’-TCG GTG TTA GAG CCT GTA AC -3’ (Li and

Wrange, 1995) (starting at the 28th bp, counting from the cleavage site of EcoR I) on the left-hand side (5’) of the cERE, 5’AGGTCActgTGACCT 3’. The pink shaded nucleotides represent the AT-rich region that are constrained so that the minor groove

faces toward the histones and the major groove is directed and exposed to the outside.

Green shaded nucleotides are the GC-rich regions that have their major groove facing inward toward the histone core. The symbol (•) depicts the center of ERE and (*) depicts the center of the DNA fragment produced by EcoR I and Hind III double digest.

7. Isolation of 161 bp DNA from plasmid

7.1 Preparation of DNA labeled on one strand

The amount of DNA was quantified using Nanodrop spectrophotometer assuming

A260 =1 for 50 μg/mL of DNA.

About 500 µg of plasmid DNA was taken and digested with EcoRI to produce

linear DNA. The enzyme was inactivated by heating at 68 oC for 20 minutes. One percent

agarose gel was run to verify the complete digestion of the plasmid DNA. The DNA was

dephosphorylated with 4 µL of 5 U/µL Antarctic phosphatase (AP) (NEB Cat # M0289S)

at 37 oC for 30 minutes, supplied with a buffer. The reaction was stopped by adding 7 µL

0.5 M EDTA (final concentration of 15 mM), 5.5 µL 20 % SDS (final concentration 0.5

%). The mixture was then treated with 2 µL proteinase K (Sigma, cat # P2308) (10

mg/mL) and incubated at 56 oC for an hour. The DNA was then extracted with phenol-

chloroform. The DNA was ethanol precipitated and stored in TE buffer at -20 oC.

7.2 Labeling at a single end of double stranded DNA

Two hundred micrograms (5µL) of the plasmid DNA was aliquoted in the 1.5 mL

Eppendorf tube. A 16 µL of dH20, 2.5 µL of 10X Opti-kinase buffer, 2.5 µL γ-(32P)-ATP

(6000 mC/mmol) and 1 µL of Opti-kinase enzyme was added to it, mixed well by gently

swirling with the pipette tip, and incubated at 37 oC water bath for 30 minutes. The reaction was stopped by the heating at 68 oC for 30 minutes. The labeled DNA was

separated from γ-(32P)-ATP by gel permeation chromatography using a spin column

containing G-50 Sephadex swollen in Millipore water. The DNA was ethanol precipitated

and dissolved in TE buffer. 38

The linear DNA was cut a second time with Hind III to produce 161 bp DNA.

The 161 bp DNA was separated from the rest of the plasmid DNA on a 6% polyacrylamide gel.

7.3 Separation of 161 bp DNA fragment from plasmid DNA

The DNA from the double digest was then loaded onto a 6% polyacrylamide gel and run for 40 minutes at 80 volts. The gel was stained with methylene blue until bands are clearly visible. This was followed by destaining in water. The DNA bands corresponding to the 161 bp (the only band in the middle of the gel) DNA was cut out of the gel with a razor.

7.4 Elution of 161 bp DNA fragment from polyacrylamide gel

The gel slice containing the 161 bp DNA fragment was transferred to a 1.5 mL

Eppendorf tube, and minced with a razor blade. A one mL of gel elution buffer (0.5 M ammonium acetate, 10 mM Mg acetate, 1 mM EDTA) was added to the gel and vortexed vigorously. The tube was incubated at 55 oC for one hour, centrifuged at 10,000 rpm and the supernatant transferred into a new 1.5 mL Eppendorf tube. An additional 200 µL of gel elution buffer was added to the gel, vortexed, microcentrifuged, and the supernatants combined. One tenth volume of 3 M sodium acetate and two volumes ice cold absolute ethanol was added to precipitate DNA for at least 4 hrs at -80 oC. The pellet was carefully washed with 100µL of ice cold 70 % ethanol, air dried, and dissolved in TE buffer. 39

8. Measurement of specific activity of DNA by scintillation counter

A sample of 100 ng 161 bp DNA was labeled with 32P-γ-ATP. The labeled DNA

was separated from 32P-γ-ATP by gel permeation chromatin using a 1 mL Sephadex G50

in a column. One µL of the labeled DNA was transferred onto a Skatron filter consisting

of a membrane filter on top and a roll of absorbent paper located beneath the filter. An

aliquot of 25 µL of ice cold 10% TCA was applied to the filter to precipitate the labeled

DNA. The DNA fragment was precipitated on the membrane and separated from the free

-32P-ATP which passed through the filter and was absorbed by the paper. The TCA

precipitation was repeated three times, and then the filter removed and transferred to a 1.5

mL scintillation vial and 1 mL of liquid scintillation solution (ECOLUME) was added to

measure the radioactivity on Beckman-LS-133 Scintillation System. The specific activity

of the DNA was determined, using the reading from the Beckman-LS-133 Scintillation

System.

A typical experimental run would have the activity of 1 µL (10 ng) aliquot of this

DNA at 66,891 cpm, which is equivalent to 9.18 * 10-14 mole DNA. Therefore, the specific activity of the DNA was 7.29 * 1017 cpm / mole DNA. In a typical reaction, 5

µL of nucleosomes was incubated in 10 µL reaction volume. The activity of 1 µL aliquot

of this nucleosomal DNA was 4209 cpm. Therefore, the concentration of nucleosomes in

a reaction was 2.85 * 10-9 M. There is a 100 fold excess of HMGB1 to nucleosomes,

because nucleosomes were incubated with 400 nM HMGB1.

9. Preparation of rotationally phase and translationally positioned nucleosomes and tailless nucleosomes containing cERE by salt dilution method (Li and Wrang, 1995).

Labeled DNA (ca. 50 ng; 10µL and 100,000 cpm ) was lyophilized in a Speed-

Vac without application of heat. To this was added 8 µL of 5 M NaCl, 4 µL of 10X reconstitution buffer (RC) (150 mM Tris-HCl (pH 7.5), 2 mM EDTA, 2 mM PMSF) and

28 µL of donor oligonucleosomes. The mixture was incubated at 37 oC for 30 minutes

and was followed by stepwise dilution during 4 hours at room temperature with 1X RC

buffer. The dilution protocol involved 5 µL of 1X RC buffer added to the sample 6 times

at 15 minutes interval, followed by the 6 additions of 10 µL of 1X RC buffer every 10

minutes. Finally, additions of 20 µL of 1X RC buffer were added every 10 minutes to

arrive at a final volume of 300 µL and the final concentration of the buffer was NaCl is

0.133 M.

The radiolabelled DNA incorporated into nucleosomes was then separated from

the large unlabelled DNA, free labeled DNA, donor oligonucleosomes and free histones

by sucrose gradient centrifugation in 5-30% TE/sucrose gradient. This was spun in

SW55Ti rotor at 36,000 rpm for 16 hours at 4 oC. Fractions of about 250 µL were

collected with the Density Gradient Fractionator by puncturing the bottom of the tube and

introducing a 50% sucrose solution from the bottom. The flow rate was 0.35 mL/min and

the samples were run on a 4% polyacrylamide gel to monitor the distribution of free

DNA and nucleosomal DNA (see results section, figure 16). 41

10. Preparation of sucrose gradient

Five, thirty and fifty per cent sucrose (w/v) solution were made in TE buffer (10

mM Tris-HCl pH 7.5, 1 mM EDTA) and autoclaved. The 5-30% sucrose gradient was prepared with a gradient maker (2.5 mL of each of 5 and 30% sucrose solution). A total of 5 mL were mixed into centrifuge tubes (Beckman Cat # 344057) for SWTi55 rotor

(2.5 mL of each of 5 and 30% sucrose). The higher sucrose solution was placed in the

right column of the gradient maker when making the gradient. The gradients were then left in the refrigerator at 4 oC to diffuse to linearity for about 4 hours. The nucleosomes

mix (300 µL) was loaded carefully on to the top of the gradient. The centrifuge tubes

were placed in the various centrifuge buckets and the whole setup balanced (with a

weighing balance) before centrifugation. (If there tubes are different in weight, adjust the volume with the 1X reconstitution buffer used in the salt dilution of nucleosomes).

11. Fractionation of nucleosomes

The samples were centrifuged at 35,000 rpm for 16 hours at 4 oC in a SW55Ti

rotor. After centrifugation, the gradient was fractionated by pushing 50% sucrose solution

up the centrifuge tube. The syringe of the Density Gradient Fractionator (ISCO, Model

185) was filled with 50% sucrose and the pump adjusted so the sucrose begins to flow

from the needles. Place the centrifuge tubes on top of the needle and push the whole setup upwards. Fit the tube tightly onto the cap on top and fractionate the gradient.

Fractions of about 250 µL were collected after poking the bottom of the tube and the 50%

sucrose solution to push up the gradient fractions. The flow rate was 0.35 mL/min. The fraction were collected into 20 tubes (1.5 mL Eppindorf tubes), on ice. Nucleosome 42 fractions were collected in tubes that contained 5 µL of 5 ng/mL BSA so that the collection of 250 µL samples now contained 0.1 µg/mL BSA. The samples were stored at

20 oC.

12. Preparation of HMGB1-remodeled nucleosomes

The fraction that contained nucleosomes was used to make the modified nucleosomes. Two hundred and eighty eight µL of the nucleosome (ca. 700 nM) was incubated with HMGB1 (12 µL of 1000 ng/µL stock HMGB1 to a final concentration

1600 nM, final volume of mixture was 300 µL). The mixture was incubated on ice for an hour and loaded onto a sucrose gradient as before. The samples were centrifuged at

35,000 rpm for 16 hours at 4 oC in a SW55Ti rotor (5 mL tubes). Fractions of about 100

µL were collected after poking the bottom of the tube and the 50% sucrose solution to push up the gradient fractions. The flow rate was 0.35 mL/min. Samples were run on a

4% polyacrylamide gel to monitor the distribution of nucleosomes (see results section, figure 27).

13. Sample preparation of HMGB1-remodeled nucleosomes

The reactions were performed in a 500 µL Eppendorf tube maintained at 4 oC. In a typical 10 µL reaction volume of nucleosomes with varying concentration of HMGB1,

8 µL of nucleosomes in TE/sucrose (18%) buffer was added to the 500 µL Eppendorf tube. To it, varying amounts of 1000 ng/µL (40 µM) HMGB1 was added and final volume of 10 µL was attained. The HMGB1 was diluted in dH2O to concentrations, such that the addition of 2 µL to a final volume of 10 µL results in final concentrations of 0, 43

400, 800 or 1600 nM. To achieve a final HMGB1 concentration of 1600 nM in a 10 µL reaction, 5 µL of the 1000 ng/µL stock was diluted to 25 µL, and 2 µL of this dilution added to 8 µL of nucleosomes. The reaction mixture was incubated on ice for 1 hour.

After incubation, the sample was loaded in the pre-electrophoresed 4% native polyacrylamide gel.

Table 3. Incubation of nucleosomes with increasing HMGB1 concentration.

Initial Nucleosome HMGB1 (µL) Buffer (µL) Final

[HMGB1] nM added (µL) [HMGB1] nM

0 8 0 2 0

2000 8 2 0 400

4000 8 2 0 800

8000 8 2 0 1600

The table shows the amount of HMGB1 added to nucleosomes, and the final concentration of HMGB1 in the reactions. 44

14. ER binding to nucleosomes

For binding studies of ER binding to nucleosomes, different concentrations of ER

were made with dilution buffer (80 mM KCl, 10% glycerol, 15 mM Tris-HCl (pH 8.0),

0.2 mM EDTA, and 0.4 mM DTT), 2 ng/µL poly (dI-dC) and 0.1 mg/mL bovine serum

albumin (BSA). The protein stock solutions ER and HMGB1 were diluted with dilution

buffer to nanomolar (nM) concentrations to be used in reactions. In a typical reaction, 5

µL of nucleosomes was incubated with 5 µL of proteins. The final reaction concentration of the various components is half the original concentration.

Five microliters of the stock HMGB1 (1000 ng/µL) were diluted to a final volume of 100 µL with ER dilution buffer, and 2µL added to the reaction tubes (see below). The final HMGB1 concentration was 10 ng/ µL (2µL in a total volume of 10µL) which is equivalent to 400 nM. HMGB1 levels were kept constant in the reactions. To prepare a

reaction that contains varying concentration of ER, the ER stock was diluted with ER

dilution buffer. For example, to prepare 0, 20, 40 and 80 nM ER reactions, 10 µL of the

stock ER (2016 nM) was diluted to a final volume of 75 µL, resulting in a final ER concentration of 268.8 nM. Nucleosomes were then incubated with 3µL of the 268.8 nM

ER resulting in a final ER concentration of 80 nM. The table below illustrates how to make various dilutions to make a final reaction volume of 10 µL. 45

Table 4. ER binding reactions to nucleosomes in the presence of 400 nM HMGB1.

Initial [ER] ER added HMGB1 Buffer (µL) Nucleosomes Final [ER]

nM (µL) (µL) (µL) nM

0 0 2 3 5 0

67.2 3 2 0 5 20

134.4 3 2 0 5 40

268.8 3 2 0 5 80

The table shows the amount of ER and HMGB1 added in ER binding to nucleosomes

(350 nM), and the final concentration of ER in the reactions.

15. DNase I digestion of nucleosomes and modified nucleosomes

15.1 DNase I 10 bp pattern

The DNase I 10 bp patterns is an indication of the rotational phasing of the

nucleosome. DNase I enzyme cuts in the minor groove of nucleosomal DNA every 10 bp.

The nucleosomes reactions were performed in a total of 200 µL at room temperature. A

tenth of the DNase I dilution buffer (100 mM Tris-HCl (pH 7.5), 20 mM MgCl2 and 5

mM CaCl2) was added to the nucleosomes. The final buffer concentrations were 10 mM

Tris-HCl (pH 7.5), 2 mM MgCl2 and 0.5 mM CaCl2. The nucleosomes were digested

with 0.04 U of DNase I. The reaction was stopped at different time interval (30, 60, 90 &

120 sec) by removing 50 µL into 5 µL of stop solution (200 ng/mL tRNA, 0.5% SDS and 46

0.3 mM Na acetate, pH 5). The fractions were treated with a final concentration of 1

µg/mL proteinase K at 55 oC for 30 minutes. The fractions were the extracted with

phenol/chloroform and chloroform alone (2x for each). Three volumes of 95% ethanol

was added and stored at -80 oC for 1 hour. The tube was then centrifuged at maximum speed on the microfuge for 10 minutes to pellet the DNA. A 250 µL of 0.3 M sodium acetate and 750 µL of ethanol was added to the pellet and stored at -80 oC for 15 minutes.

The tube was then centrifuged at 10,000 rpms and the supernatant carefully removed.

The DNA precipitate was washed with 70% ethanol. The pellet was the dissolved in 20

µL of TE and lyophilized. The cpm were counted with BC 2000 counter and the

radioactivity adjusted so as to have the same number of counts in each well.

15.2 DNase I footprint

The location of the binding site for a sequence specific transcription factor can be

identified by a DNase I footprint. To determine a DNase I footprint for ER bound to the

cERE in the nucleosome, the normal procedure for EMSA is followed, with a few

modifications. The reactions were performed as in the EMSA in a total of 200 µL.

Nucleosomes (100 µL) was incubated with increasing concentration of ER in the

presence of 400 nM HMGB1. The ER and HMGB1 proteins were diluted with ER

dilution buffer (80 mM KCl, 15 mM Tris-HCl (pH 8.0), 0.2 mM EDTA, 0.4 mM DTT and 10% glycerol); and 2 ng/µL poly (dI-dC) with 0.1 mg/mL BSA. The reaction was incubated on ice for 30 minutes and at room temperature for 5 minutes. A tenth volume

of DNase I buffer was added to the reaction and the steps for the DNase I followed. The 47

final concentrations of the buffer components were 8 mM KCl, 10 mM Tris-HCl, 0.2 mM

EDTA, 0.04 mM DTT and 5% glycerol), 1 ng/µL poly (dI-dC).

16. Exonuclease III digestion of nucleosomes

Exonuclease III (Exo III) is a probe that can determine the extent of exposure of

DNA from the end of the nucleosome. Exo III (200 U/ µL, Fisher Scientific Cat #

BP3213-1) digestion of nucleosomes was performed in a total of 200 µL at room

temperature. A tenth volume of the Exo III dilution buffer (the DNase I buffer was used)

was added to the nucleosomes. The nucleosomes were digested with 200 U of Exo III.

The reaction was stopped at different time interval (0, 1, 2 & 4 minutes) by removing 50

µL into 5 µL of stop solution (200 ng/mL tRNA, 0.5% SDS and 0.3 mM Na acetate). The fractions were treated with a final concentration of 1 µg/mL proteinase K at 55 oC for 30

minutes. The fractions were the extracted with phenol/chloroform and chloroform alone

(2x for each). Three volumes of 95% ethanol was added and stored at -80 oC for 1 hour.

The tube was then centrifuged at maximum speed on the microfuge for 10 minutes to

pellet the DNA. A 250 µL of 0.3 M sodium acetate (to redissolve the pellet) and 750 µL

of ethanol was added to the pellet and stored at -80 oC for 15 minutes. The tube was then

centrifuged at 10,000 rpms and the supernatant carefully removed. The DNA was

washed with 70% ethanol. The pellet was then dissolved in 20 µL of TE and lyophilized.

The cpm was counted with BC 2000 counter and the radioactivity adjusted so as to have

the same number of counts in each well.

17. Ava I digestion on nucleosomes and modified nucleosomes

Ava I restriction enzyme digest is used to test the accessibility of nucleosomes.

The Ava I (50 U/ µL) (New England Biolabs, Cat # R0152T) restriction enzyme digestion

of nucleosomes and modified nucleosomes were performed in a total of 200 µL reaction

volumes using 100U of Ava I, at 37 oC. A tenth volume of the 10X Ava I buffer (50 mM

potassium acetate, 20 mM Tris-acetate, 10 mM magnesium acetate and 1 mM DTT) was

added to the nucleosomes. The final buffer concentration was 5 mM potassium acetate, 2

mM Tris-acetate, 1 mM magnesium acetate and 0.1 mM DTT. The reaction was stopped

after 0, 5, 10, 15 and 30 minutes by removing 40 µL into 10 µL of stop solution (20 mM

Tris-HCl, 1 mg/mL tRNA, 10% SDS and 50 mM EDTA). The fractions were treated with

a final concentration of 1 µg/mL proteinase K at 55 oC for 30 minutes. The fractions were

the extracted with phenol/chloroform and chloroform alone (2x for each). Three volumes

of 95% ethanol was added and stored at -80 oC for 1 hour. The tube was then centrifuged

at 10,000 rpms and the supernatant carefully removed. The DNA precipitated was

washed with 70% ethanol. The pellet was then dissolved in 20 µL of TE and lyophilized.

The lyophilized fractions were normalized with denaturing gel loading buffer (80 % (v/v) formamide, 50 mM Tris-Borate, 1 mM EDTA, 0.1 % (w/v) xylene cyanol, 0.1 % bromophenol blue), and run on 8% denaturing gel.

18. Preparation of DNA A/G ladder to define position on DNA

DNA that was labeled at the EcoRI end was chilled to 4 oC and 10 µL was pipetted into an Eppendorf tube. The volume was adjusted to 20 µL with chilled

Millipore water and treated with 5 µL of formic acid(Fisher, Cat # 7732-185) that was 49

diluted 1 in 10 with Millipore water. The tube was then incubated at 37 oC for 5 minutes.

The reaction was stopped with the addition of 250 µL of HZ stop solution (0.3 M EDTA,

25 µg/mL tRNA and 0.3 M sodium acetate), 750 µL of 95% ethanol added and incubating at -80 oC for 15 minutes. The tube was then centrifuged at maximum speed

on the microfuge for 10 minutes to pellet the DNA. A 250 µL of 0.3 M sodium acetate

(to re-dissolve the pellet) and 750 µL of ethanol was added and stored at -80 oC for 15

minutes. The mixture was again centrifuged at maximum speed and the supernatant

carefully removed. The pellet was then washed with 100 µL of 70 % ethanol and

vacuum dried. The dried pellet was then treated with 100 µL piperidine (Aldrich, Cat #

104094) that was diluted 1 in 10 with Millipore water, and heated at 90 oC for 30 minutes

in a heating block. The mixture was lyophilized in a Speed Vac. The pellet was washed

twice by re-dissolving in 10 µL of water and lyophilizing. Finally the pellet was

dissolved in 10 µL of loading buffer (80 % (v/v) formamide, 50 mM Tris-Borate (pH

8.3), 1 mM EDTA, 0.1 % (w/v) xylene cyanol, 0.1 % bromophenol blue). The mixture

was then vortexed and heated on a heating block for 1 minute at 90 oC and chilled on ice.

This will be used to produce an A/G ladder for a sequencing gel and be used as a nucleotide marker in DNase I and Exo III digestions.

19. Electrophoretic Mobility Shift Assay (EMSA)

19.1 Gel preparation

A 4% native polyacrylamide gel was used for the EMSA studies. To prepare a

4% polyacrylamide gel, a 3.5 mL aliquot of 10X TBE buffer (900 mM Tris-HCl, 900 mM boric acid, and 20 mM EDTA), 13.4 mL 30% (w/v) polyacrylamide (29:1, 50

polyacrylamide: bisacrylamide), 1 mL of 5% IGEPAL and made to 100 mL with

Millipore water were dispensed into a 500 mL flask and degassed for 15 minutes. After

degassing, a 500 µL of 10% (w/v) ammonium persulfate and a 100 µL of TEMED were added and mixed well. The mixture was carefully poured into a pre-assembled glass

plates (19.6 x 19 cm) and left to polymerize.

A denaturating 8% polyacrylamide sequencing gel was prepared in 1X TBE

buffer. A 5 mL aliquot of 10X TBE buffer (900 mM Tris-HCl, 900 mM boric acid, and

20 mM EDTA), a 16 mL 25% (w/v) polyacrylamide (19:1, polyacrylamide:

bisacrylamide) and a 21 g of urea (Fisher Scientific, Cat # U17-212) were dispensed into

a 100 mL beaker. The mixture was heated gently while stirring to dissolve the urea. The volume was then adjusted with dH2O to 50 mL and filtered through 3 M Whatman filter paper using a water aspirator for a vacuum suction. The mixture was then degassed for 15 minutes and cooled on ice to slow down the polymerization process. Two hundred and

fifty microliters of 10% ammonium persulfate and 50 µL of TEMED were then added to

it, mixed well and the gel mixture poured into the pre-assembled glass plates. The gels

are pre-electrophoresed until the temperature increased to 45 oC (determined by

temperature strips) and the current decreased to half of its initial value before loading the

samples.

19.2 DNA gel electrophoresis

The 4% polyacrylamide gel was assembled in the electrophoresis apparatus

(Vertical Gel Electrophoresis System, Model V16, Cat. # 21070, Whatman, Inc). The

upper and lower chambers were filled with 0.35X TBE. The EMSA gels were pre-

electrophoresed at 100V for one hour or until the current dropped to half its starting 51

value. The voltage was increased to 200V and then the samples were loaded into the

wells and electrophoresed typically for 2 hours at 4 oC.

The 8% sequencing gel was prepared in 20 X 40 cm glass plates, one of which is

siliconized on the surface. The upper and lower chambers were filled with 1X TBE. The

gel was pre-electrophoresed at 1500 V until the temperature increases to 45 oC before

loading the samples. The samples were electrophoresed for 3 hours.

19.3 Gel drying

After the electrophoresis was complete, the gel cassette was removed from the apparatus and the two glass plates were pried apart. The fragile gel tended to stick to one side of the glass plate. A piece of 3 M Whatman paper cut to the dimensions of the gel was placed on top of the gel, and the gel was removed from the glass plate. The gel/filter paper combination was covered with Saran Wrap and dried in a gel drying unit using a vacuum pump (Physical Science Building, Room # 216).

20. Autoradiogram

After the gel was completely dried, it was transferred to an X-Ray cassette containing an intensifying screen. An X-ray film (8” x 10”) (Merry X-Ray Corporation) was placed on the dried gel in the dark and the cassette cover was closed. The gel cassette was incubated at -80 oC for various lengths of time depending on the activity of the

radiolabeled oligonucleotide. The cassettes were removed from the -80 oC freezer and

allowed to warm up to room temperature. The film was developed in the dark by

immersing and gently agitating the film for 5 minutes in developer solution (Kodak GBX 52 developer, Cat # 1900984). The film was rinsed in distilled water and finally it was placed in fixer solution (Kodak GBX fixer, Cat # 1902485) for 2 minutes. The film was then rinsed once more with the distilled water and then hung up to dry.

21. SDS-PAGE of proteins

21.1 Gel preparation

The BioRad protein mini-gel apparatus was used for both casting and running gels. An appropriate amount of acrylamide solution was used to obtain a desired final percentage of acrylamide in the separating layer. Typically, an 18% polyacrylamide gel was used. A 15 mL separating gel solution was prepared. In a 25 mL Erlenmeyer flask, a

6 mL of 30% (29:1, acrylamide: bisacrylamide), 2.5 mL of 1.5M Tris-HCl, pH 8.8, 100

µL of 10% SDS and 1.4 mL of Millipore water was added and mixed well. This solution was degassed for 15 minutes and 50 µL of 10% APS and 10 µL TEMED was added to it and mixed well. The polymerizing solution was immediately poured between two glass plated up to the level of 1 cm below where the comb in the stacking layer would be. A layer of water saturated isobutanol was poured on top of the gel while polymerization was completed (in about 25-45 minutes). A solution of 4% acrylamide was used to prepare the stacking layer. In a 25 mL Erlenmeyer flask, 650 µL of 30% (29:1 acrylamide: bisacrylamide), 1.25 mL of 0.5 M Tris-HCl, pH 6.8, 50 µL of 10% SDS and

3.05 mL of dH2O was mixed well. The stacking gel mixture was degassed for 15 minutes and 25 µL of 10% APS and 5 µL of TEMED were added to it and mixed well. The polymerizing layer was poured on top of the already polymerized separating gel. The 53

comb was immediately placed between the two glass plated of the gel to form the sample

wells.

21.2 Sample preparation for proteins

A calculated volume of Blue juice (10X SDS loading buffer (250 mM Tris-HCl

pH 6.8, 10% SDS, 30% glycerol, 5% β-mercaptoethanol, 0.02% bromophenol blue) was

added to the sample to attain 1X final concentration. The samples were mixed well and

briefly centrifuged at high speed in a microcentrifuge. The samples were heated in a

boiling water bath for 5 minutes and spun down. The sample was then loaded on the gel.

21.3 SDS-PAGE

After polymerization was complete, the comb in the stacking layer was removed

and the wells were thoroughly washed with distilled water. The BioRad gel cassette was

assembled and then fitted onto the BioRad mini gel electrophoresis unit. Pre-

electrophoresis was performed at 150V for 20-30 minutes. Samples prepared in a final concentration of 1X loading buffer (Blue juice) was loaded into the wells and the gel was electrophoresed at 200V in electrophoresis buffer (25 mM Tris-HCl, 192 mM glycine,

0.1% (w/v) SDS). The blue dye gave an indication as to when the gel should be stopped.

Once the dye ran off the gel, the electrophoresis was stopped.

21.4 Gel staining

After electrophoresis was complete, the gel was removed from the glass plate sandwich and placed in 50% ethanol, 10% acetic acid and heated in a microwave for 1 minute. The gel was then placed on a shaker for 5 minutes at room temperature, during 54

which the gel shrinks. The gel was resuspended in 5% ethanol, 7.5% acetic acid and 0.2%

Coomassie blue R-250. The container was heated in the microwave for 30 seconds and left on the shaker for 30 minutes at room temperature. The bands corresponding to proteins become visible after this time.

21.5 Gel preservation

The gel was placed in soaking solution (20% (v/v) ethanol and 10% (v/v) glycerol) in order to avoid cracks in the gel when they were dried. The cellophane used to dry the gel was submerged in distilled water for 15 minutes. After the soaking process was complete, one sheet of cellophane paper was smoothed across a plexi-glass plate.

Then the gel was placed on top of the cellophane sheet making sure that there were no air bubbles. Finally the other piece of cellophane was placed on top of the gel and smoothed out so that there were no air bubbles. A small plexi-glass frame was placed over the last piece of cellophane, and it was secured by binder clips. The gel was allowed to dry for several days at room temperature which it was placed in the lab notebook or scanned.

22. Quantification of radioactivity to determine KD values

The polyacrylamide gels were exposed to a phosphoimager screen for 1-3 days

and the screen scanned using the Storm Scanner. The bands were then quantified with

Image Quant software. The intensity of the bands is directly proportional to the amount

of radioactivity in the band. The KD values were then quantitatively determined from these readings. The concentration of labeled DNA is the same in all the lanes.

The dissociation of the receptor from DNA can be represented as: 55

(Protein/DNA)complex = Proteinfree + DNAfree

KD= [Proteinfree] [DNAfree] / [Protein/DNA]

The point at which 50 % of the labeled DNA is complexed with the protein is taken as the

KD value.

When 50% of the protein is bound to DNA, [DNAfree] = [Protein/DNA]

 KD = [Proteinfree]

Percentage complex formed was determined from the intensities of the complex and free

DNA measured by phosphoimager as

% complexation = [complex] / [complex + DNA]

A graph of percentage complex versus protein concentration was prepared on a Sigma

Plot program and the KD was determined from the plot at the 50 % complexation.

Table 5. Final concentration of each component of the reaction buffer

Buffer Components* Final Conc. ER dilution buffer Tris-HCl 10 mM KCl 40 mM (80 mM KCl, 10% glycerol, 15 mM Tris- EDTA 0.6 mM HCl pH 8.0, 0.2 mM EDTA, 0.4 mM DTT, DTT 0.2 mM 100 ng/µL BSA, 2 ng/µL poly (dI-dC)) BSA 50 ng/µL poly (dI-dC) 1 ng/µL ER dilution buffer minus poly (dI-dC) Tris-HCl 10 mM KCl 40 mM (80 mM KCl, 10% glycerol, 15 mM Tris- EDTA 0.6 mM HCl pH 8, 0.2 mM EDTA, 0.4 mM DTT, DTT 0.2 mM 100 ng/L BSA) BSA 50 ng/µL Ava I digestion MgAc 1 mM 2 50 mM potassium acetate, 20 mM Tris- DTT 0.1 mM acetate, 10 mM magnesium acetate and Tris-acetate 2 mM 1 mM DTT. KAc 5 mM

DNase I Tris-HCl 10 mM

100 mM Tris-HCl (pH 7.5), 20 mM MgCl2 MgCl2 4 mM and 5 mM CaCl 2 CaCl 0.5 mM 2 Exo III Tris-acetate 3.3 mM

MgAc 1.0 mM 33 mM Tris- acetate, pH 7.8, 66 mM KAc, 2 KAc 6.6 mM 10 mM MgAc , 0.5 mM DTT 2 2 DTT 0.05 mM

*Note: The reaction mixture was prepared by mixing one part ER dilution buffer and one part nucleosomes in 18% sucrose/TE buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA). Therefore, the final concentration of sucrose is 9% and glycerol is 5% in all the reactions **Exo III buffer converts N’/N” to canonical nucleosomes and was therefore, not used in Exo III digestion. All Exo III digestions were done in DNase I buffer which kept the Exo III activity and did not convert any of N’/N” to canonical form.

CHAPTER 3: RESULTS

1. Isolation and purification of HMGB1 and HMGB2 proteins from calf thymus

Our laboratory has shown that HMGB1 enhances the binding of ER to cERE on 33 bp DNA (Das et al, 2004). We set out to expand this previous study to ER binding to cEREs in rotationally phased and translationally positioned nucleosomes.

HMGB1 and HMGB2 proteins were isolated from calf thymus as described in (See chapter 2), (Das, 2002). The proteins were purified by HPLC using Mono Q 5/5 anion exchange column, with a linear salt gradient (0.2-0.8 M NaCl). The elution time for

HMGB2 and HMGB1 were 13.03 and 14.05 respectively, on an18 min linear salt gradient.

The purity of the fractions was verified with 18% SDS-PAGE as shown in figure 11, with only a single band observed for each HMGB1 and HMGB2 at ca 25 kDa, which is the appropriate molecular weight for the HMGB proteins (Johns, 1982). However, the two proteins can be distinguished since HMGB2 exhibits a slightly greater mobility compared to HMGB1 as shown in the figure 11 (Johns, 1982). 58

Figure 11. An 18% SDS-PAGE of purified HMGB1 and HMGB2. Lane 1, molecular weight markers; lane 2, HMGB1 fraction; lane 3, HMGB2 fraction. The molecular weight markers were 15, 20, 25, 37, 50, 75, 100, 150 and 200 kDa.

2. Preparation and isolation of oligonucleosomes

2.1 Micrococcal nuclease digestion

To prepare nucleosomes, a source of donor core histones was needed. Chromatin

from chicken erythrocyte (CE) was selected as the source. The nuclei from chicken erythrocytes (5 mg/mL DNA; assuming A260 = 50 µg/mL in a total volume of 5 mL) was

digested with 75U of micrococcal nuclease (MN) for different time intervals (5-25

minutes) and the reaction stopped by addition of EDTA to 2.2 mM (see chapter 2). Figure

12 shows that a 5 minute digestion results in a range of oligonucleosomes, primarily with

DNA sizes greater than 1000 bp DNA or 5 nucleosomes. Digestion of the chromatin for 25

minutes resulted in the formation of nucleosomes that contained about 150-200 bp of DNA

(core particle) as shown in figure 12, lane 6. Since we were interested in using oligonucleosomes with DNA fragments much larger than 200 bps, subsequent digestions

with MN were performed for less than 5 minutes.

2.2 Gel permeation chromatography of MN digestion of chromatin

Gel permeation chromatography separates proteins and particles according to size

as presented in chapter 2. Using a Sepharose CL-4B column and 0.65M NaCl buffer, H1

protein and non-histone proteins were dissociated and separated from oligonucleosomes

that contained only the core histones. These oligonucleosomes were then used as the donor

of core histones in the preparation of radiolabeled nucleosomes (Li and Wrange, 1995).

Oligonucleosomes containing only the core histone proteins intact were eluted on

the Sepharose CL-4B column with 0.65 M NaCl buffer from the digested chicken

erythrocyte (CE) chromatin. The purity of the core histones within the oligonucleosomes 60 was verified by 18% SDS-PAGE. Figure 13 shows only four bands representing the core histone proteins from the fractions (H2A, 13.9 kDa; H2B, 13.7 kDa; H3, 15 kDa and H4,

11 kDa), with no evidence of the presence of histone H1, (21 kDa). 61

Figure 12. Micrococcal nuclease digestion of CE chromatin. Lane 1, 100 bp ladder (100,

200, 300, 400, 500, 600, 700, 800, 900, 1000, 1100, 1200, 1500, 1600, 1800 and 2000);

Lane 2-6, chromatin digested with 75U micrococcal nuclease for 5, 10, 15, 20 and 25 minutes, respectively.

Figure 13. SDS-PAGE of core histone proteins in oligonucleosomes. Lanes 1, molecular weight markers; lane 2, micrococcal nuclease digested chromatin extract; and lane 3, fractionated oligonucleosomes. The results show only four bands representing the core histone proteins (H2A, 13.9 kDa; H2B, 13.7 kDa; H3, 15 kDa and H4, 11 kDa). The bands for core histones run slightly different from the molecular weight markers because histone proteins are highly charged. H1 with molecular weight of 21 kDa appears at ca. 27 kDa in lane 2. 63

3. Preparation of nucleosomes

3.1 Diagram of 2E2, 3E1 and 4E0

The DNA constructs (2E2, 3E1 and 4E0) were prepared by Dr Ron Peterson (details in chapter 2). All constructs contain 4 nucleosome positioning sequences (NPS) (TCG GTG

TTA GAG CCT GTA AC; Wrange, 1993). The 2E2 has the cERE centered at the dyad axis of the DNA, with two NPS on each side of the cERE. The 3E1 has the cERE centered at 20 bp from the dyad axis and contains three NPS on the EcoRI side of the cERE and one on the Hind III side of the cERE. The 4E0 contains cERE the centered 40 bp from the dyad axis with 4 NPS on the EcoRI side of the cERE (figures 9 and 10, chapter 2). Figure 14 shows the three DNA constructs of interest. 64

Figure 14. Schematic diagram of DNA used to prepare nucleosomes. AvaI site is 98,

118 and 138 bp from the EcoRI labeled end of the DNA for 2E2, 3E1 and 4E0, respectively. The cleavage site of AvaI is 10 bp from the end of the cERE and is indicated by the arrows. The bar represents the cERE 74-88 bp for 2E2; 94-108 bp 3E1 and 114- 128 bp for 4E0. The “bricks” (although not to size) represent the blocks of nucleosome positioning sequences (NPS). The star (*) represents the DNA strand that is labeled. The dark bars at the 5’-end are the 4 bp DNA overhang due to asymmetric EcoRI and Hind III cutting. 65

3.2 Histone exchange with MN digested chromatin

We set out to extend our studies to the binding of ER to cERE within nucleosomes

and to determine whether HMGB1 influences the binding. Nucleosomes were prepared

using salt dilution method as described (chapter 2) (Li and Wrange, 1995). The 32P labeled

161 bp DNA containing the cERE at different translational positions were mixed at 1 M

NaCl buffer, with MN digested chromatin, and the high salt diluted sequentially to 0.13M.

The formed nucleosomes were separated from the unincorporated DNA on 5-30%

TE/sucrose gradient (M and M). Aliquots of 250 µL were collected by puncturing the bottom of the centrifuge tube and pushing 50% sucrose solution through the bottom of the centrifuge tube. Figure 15 shows two peaks in the sucrose gradient sedimentation profile, with fraction 5 containing unincorporated DNA and fraction 10 containing newly formed nucleosome. About a 10 µL sample of the various fractions was run on a 4% polyacrylamide gel as shown in figure 16 to determine if free DNA and DNA incorporated into nucleosomes were separable. Fractions containing nucleosomes were collected and stored at –20 oC.

Figure 15. Sucrose gradient sedimentation profile showing free DNA and nucleosomes. Individual fractions were collected from 5-30% linear sucrose gradient by pushing a 50 % sucrose solution from the bottom of the 5 mL centrifuge tube. Samples were collected (250 µL) and the counts per minute determined with the DuPont BC 2000 counter. Fractions 5 contain labeled DNA, while fractions 10 and 11 contain nucleosomal

DNA. The fractions were stored at -20 oC. 67

Figure 16. Sucrose gradient centrifugation fractions analyzed on polyacrylamide gel.

About a 10 µL sample of the various fractions was run on a 4% polyacrylamide gel. The bands corresponding to nucleosome were not contaminated with free DNA.

4. Binding of ER to DNA

To ensure that ER binds to ERE in the DNA, binding studies were performed with

ER. Figure 17 shows the ER binding profile on 4E0. The KD value calculated was about 5 nM. This was consistent with previous results in our laboratory which showed ER binds to

161 bp DNA with a KD of 3 nM (Sarpong, 2006). Figure 18 shows the binding profile of three separate experiments of ER binding to 4E0. The KD was also determined for 3E1 and

2E2 and was determined to be independent of the translational positioning of the cERE. 69

Figure 17. ER binding profile to 4E0 DNA. DNA was incubated with increasing concentrations of ER on ice for 30 minutes and run on 4% polyacrylamide gel for 2 hours.

Lane 1-6 contained ER concentrations 0, 1, 2, 4, 6 and 8 nM, respectively.

30 % Complex 20

0 0.1 1 10 ER (nM)

Figure 18. Binding profile of ER binding to 4E0. The intensity of the bands on the gel was determined using a phosphoimager and % complex plotted from three independent gels, using the Origin 6.2 software. ER concentrations were 0, 1, 2, 4, 6 and 8 nM.

5. DNase I footprint on the free DNA

A DNase I footprint was used to determine if the binding observed for ER binding to DNA was sequence specific. All DNA used in this experiment was labeled at the EcoRI

end. Figure 19 shows a footprint of ER binding on the free DNA (2E2 and 4E0) as the

concentration of ER was increased from 0 to 40 nM. Figure 19 shows that there was a

decrease in cutting at 74-88 bp and 114-128 bp for 2E2 and 4E0, respectively, which were

the locations of the cERE. Therefore, this indicates that ER binds to cERE specifically and

reduces the accessibility of the site to DNase I cutting. 72

Figure 19. DNase I footprint on DNA containing either 2E2 and 4E0. DNA was incubated with increasing concentrations of ER on ice for 30 minutes. The tubes were then kept at room temperature for 5 minutes and digested with 0.04 U DNase I. The reaction was stopped after 60 seconds by adding EDTA to a final concentration of 25 mM. The resulting DNA was purified and run on 8% denaturing gel for 2.5 hours. Lane 1, A/G ladder; Lane 2-6 and 7-11 contained ER concentrations 0, 5, 10, 20 and 40 nM, respectively. The bars on the side of the figure denote the location of the cERE; 74-88 bp for 2E2 and 114-128 bp for 4E0, respectively. 73

6. Binding of ER to nucleosomes in the absence and presence of 400 nM HMGB1

We now examined the ER binding to cERE in a nucleosomal DNA in the absence

and presence of 400 nM HMGB1. Figure 20A shows that ER does not bind significantly to

nucleosomes in the absence of HMGB1 (perhaps a few %) but in the presence of 400 nM

HMGB1, ER binds strongly to cERE in 4E0 nucleosomes (figures 20B) and 2E2

nucleosomes (figure 22). The binding has been shown previously to be independent of the position (2E2, 3E1 or 4E0) of the ERE on the nucleosomes, with a KD of about 50 nM

(Sarpong, 2006). Figures 21 (4E0) and 23 (2E2) show the binding profile of three separate

experiments of ER binding to nucleosomes in the presence of 400 nM HMGB1, with a KD of 50 nM. The graph was plotted dividing the amount of complex formed by the total amount of DNA (see chapter 2).

Figure 20. ER binding to 4E0 nucleosomes in the absence (A) and presence of 400 nM

HMGB1 (B). Nucleosomes were incubated for 30 minutes with increasing concentration of ER on ice, and run on 4% polyacrylamide gel for 2 hours. Lanes 1-8 represents ER concentrations 0, 10, 20, 30, 40, 50, 60 and 80 nM, respectively. ER binds significantly to nucleosomes only in the presence of HMGB1.

% Complex 20

110

ER (nM)

Figure 21. Binding profile of ER binding to 4E0 nucleosomes in the presence of 400

nM HMGB1. Phosphoimager intensities of the DNA and complex bands were determined

and % complex plotted from three independent gels, using the Origin 6.2 software. ER

concentrations were 0, 10, 20, 30, 40, 50, 60 and 80 nM. The profile for ER binding to

nucleosome without HMGB1 could not be plotted because ER binding is not significant

without HMGB1. 76

Figure 22. ER binding to 2E2 nucleosomes in the presence of 400 nM HMGB1.

Nucleosomes were incubated for 30 minutes with increasing concentration of ER on ice, and run on 4% polyacrylamide gel for 2 hours. Lanes 1-8 represents ER concentrations 0,

10, 20, 30, 40, 50, 60 and 80 nM, respectively. ER binds significantly to nucleosomes only in the presence of HMGB1. 77

100

% Complex 20

1 10 100

ER [nM]

Figure 23. Binding profile of ER binding to nucleosomes in the presence of 400 nM

HMGB1. Phosphoimager intensities of DNA and complex band were determined and % complex plotted using the Origin 6.2 software. ER concentrations were 0, 10, 20, 30, 40,

50, 60 and 80 nM. The profile for ER binding to nucleosome without HMGB1 could not be plotted because ER binding is not significant without HMGB1.

7. DNase I footprint of ER/cERE in 2E2 nucleosomes

A DNase I footprint was performed on the binding of ER to nucleosomes in the presence of 400 nM HMGB1 to determine whether the binding of ER was sequence specific. Figure 24 shows a cERE footprint when 50 and 100 nM ER were reacted with 2E2 nucleosomal DNA. Figure 24 shows that there is a decrease in cutting at 74-88 bp, which is the location of the cERE, from the Eco RI end. Therefore, this indicates that ER binds to cERE and reduces the accessibility of the site to DNase I cutting. 79

Figure 24. DNase I footprint on 2E2 nucleosome. Nucleosomes were incubated on ice

for 30 minutes with 400 nM HMGB1 and increasing concentration of ER. The reactions

were then kept at room temperature for 5 minutes and digested with 0.04U DNase I. The reaction was stopped after a minute of digestion with DNase I by increasing the EDTA

concentration to 25 mM. The purified DNA was run on 8% polyacrylamide gel for 2.5

hours. Lane 1, A/G ladder; lanes 2-4 show increasing concentration of ER 0, 50 and 100

nM ER. The bar on the left side of the figure denotes the position of the cERE (74-88bp)

and the apparent DNase I footprint. 80

8. HMGB1 does not alter the rotational position of DNA within nucleosomes

I was interested as to how the HMGB1 enhances the binding of ER to cERE within

a nucleosome. DNase I can be used to define the rotational phasing of the DNA within a

nucleosome (Imbalzano et al, 1994). For a well-defined nucleosome, the radiolabeled DNA

is rotationally phased so that the DNase I cuts in the exposed minor groove every 10 bp

(with some being more intense than others). If the phasing of the DNA that is wrapped

around the core histones is altered, the 10 bp pattern would be changed, usually with the

addition of more bands, and the pattern approaching that for free DNA (Imbalzano et al,

1994).

I asked whether HMGB1 could alter the phasing of nucleosome and change the 10

bp pattern on the HMGB1-remodeled nucleosomes. Figure 25A shows that the DNase I

pattern of free DNA, in the absence or presence of 400 nM HMGB1 remains the same.

This suggests that the HMGB1 does not bind to the DNA at such a concentration and cause

a change in DNase I pattern. Figure 25B shows that the DNase I pattern in the absence or presence of 400 nM HMGB1 remained unchanged with no additional bands observed.

Therefore, 400 nM HMGB1 does not alter the rotational position of DNA in the

nucleosomes, suggesting that the interaction between DNA and the core histones is not

altered. 81

Figure 25. DNase I 10 bp profile for free DNA, nucleosomes (2E2) and nucleosomes

treated with 400 nM HMGB1. A) DNA was untreated or treated with 400 nM HMGB1, and incubated on ice for one hour. The reaction was transferred to room temperature for 5 minutes followed by DNase I digestion with 0.04U DNase I. Lane 1, DNA and lane 2,

DNA treated with 400 nM HMGB1. B) Nucleosomes were either untreated or treated with

400 nM HMGB1 and incubated on ice for one hour. The reaction was transferred to room temperature for 5 minutes followed by DNase I digestion. Lane 1, A/G ladder; lanes 2-5, nucleosomes digested for 30, 60, 90 and 120 seconds, respectively; lanes 6-9, nucleosomes treated with 400 nM HMGB1 digested for 30, 60, 90 and 120 seconds, respectively. The purified DNA was run on 8% denaturing gel for 2.5 hours. 82

9. Effect of increasing levels of HMGB1 on nucleosome mobility

To determine the effect of increasing levels of HMGB1 on nucleosome characteristics, the nucleosomes were incubated with 400, 800 or 1600 nM HMGB1 for one hour at 4 oC. The nucleosomes were then analyzed on a 4% polyacrylamide gel to determine if HMGB1 changed the mobility of the nucleosomes on electrophoretic mobility shift assay (EMSA) gel. Figure 26 shows that nucleosomes exhibit a reduced mobility as the concentration of HMGB1 was increased from 400 to 1600 nM. The mobility of the bands decreased with the addition of increasing amounts of HMGB1 in the reaction.

This suggested that HMGB1 was either stably binding to the nucleosomes and decreasing the mobility or simply altering the structure of the nucleosome. Previous work showed that HMGB1 did not form a stable component of the newly formed complex, as no supershif occurred in the position of the bands on addition of anti-HMGB1 (Sarpong,

2006). Therefore, the reduced mobility of the newly formed complex in the presence of

HMGB1 could not be due to stable HMGB1 binding to the nucleosomes (i.e., part of the

HMGB1/nucleosome complex). 83

Figure 26. Incubation of nucleosomes with increasing levels of HMGB1. The nucleosomes were incubated with increasing concentrations of HMGB1 on ice for 1 hour and run on 4% polyacrylamide gel for 2 hours. Lane 1-4 contained 0, 400, 800 and 1600 nM HMGB1, respectively. 84

10. Effect of increasing HMGB1 levels on ER binding to nucleosomal DNA

To determine the effect of increasing HMGB1 levels on ER binding, varying ER

concentrations were incubated with 200, 400 and 800 nM HMGB1. Figure 27 shows that

200 nM HMGB1 enhanced ER binding to a nucleosomes (lanes 1-6), with 400 nM

HMGB1 appearing to further increase the extent of binding (lanes 7-12). At 800 nM

HMGB1, ER also bound to cERE in nucleosomes (lanes 13-18). Therefore, it appears that

400 nM is a saturating concentration of HMGB1 that facilitates ER binding to the

nucleosome. At 1600 nM HMGB1, multiple bands for perhaps multiple complexes were

observed which may be due to also the high concentration of HMGB1 proteins binding to

the nucleosomes (data not shown). Figure 28 shows the binding profile of three separate

experiments of ER binding to nucleosomes in the presence of 200, 400 and 800 nM

HMGB1, with a KD of 47 nM. The graph was plotted dividing the amount of complex

formed by the total amount of DNA (see chapter 2). Increasing the concentration of

HMGB1 beyond 200-400 nM did not significantly enhance binding of ER to the

nucleosomes. 85

Figure 27. ER binding to nucleosomes in the presence of increasing concentration of

HMGB1. Nucleosomes were incubated for 30 minutes on ice with increasing

concentrations of ER and run on 4% polyacrylamide gel for 2 hours. Lanes 1-6, 7-12 and

13-18 contained 200, 400 and 800 nM HMGB1, respectively. Lanes 1, 7 and 13, 0 nM;

Lanes 2, 8 and 14, 10 nM; Lanes 3, 9 and 15, 20 nM; Lanes 4, 10 and 16, 40 nM; Lanes

5, 11 and 17, 60 nM; and Lanes 6, 12 and 18, 80 nM. 86

100

% Complex 20

110100 ER (nM)

Figure 28. Binding profile of ER binding to nucleosomes in the presence of 200, 400 and 800 nM HMGB1. The band intensities from phosphoimager were determined and % complex plotted from three independent gels, using the Origin 6.2 software. ER concentrations were 0, 10, 20, 40, 60 and 80 nM. The dots (●) indicate binding in the presence of 200 nM, squares (■) 400 nM, and triangle (▲) 800 nM of HMGB1.

11. Isolation of HMGB1-remodeled nucleosomes

To further characterize the nucleosomes in the lower mobility band formed in the

presence of high levels of HMGB1, we attempted to isolate them using a linear sucrose

gradient. Nucleosomes were treated with 1600 nM HMGB1 for an hour on ice and the

reaction mixture loaded onto a 5-30% TE/sucrose gradient (see chapter 2). Fractions were

collected as outlined (see chapter 2), and the nucleosome mobility was analyzed on a 4%

polyacrylamide gel.

Figure 29 shows that fractions 19-22 contained two populations of HMGB1- remodeled nucleosomes (HR-NS) which were designated N’ and N” and having a distinctly lower mobility that the canonical nucleosome (N). However, both populations of HR-NS had the same sedimentation characteristics. These N’/N” remodeled nucleosomes sedimented at 18% TE/sucrose in the 5-30% gradient. 88

Figure 29. Sedimentation profiles of remodeled nucleosomes. Nucleosomes were incubated with 1600 nM HMGB1 on ice for 1 hour and sedimented on 5-30% TE/sucrose gradient for 16 hours (See chapter 2). Aliquots from each fraction were loaded on 4% gel and electrophoresed for 2 hours at 200V. The EMSA profile shows nucleosome (N) from untreated preparations and two different bands (N’/ N”) for nucleosome altered by reaction with 1600 nM HMGB1. In the absence of HMGB1, N occurs in effectively the same fraction number as does N’/ N”.

12. Determination of the presence of core histones in nucleosomes and N’/ N”

To verify that the canonical nucleosome and the N’/N” nucleosome contained all four core histones, nucleosomes and HMGB1-remodeled nucleosomes (N’/N”) were incubated with antibodies (αH2A, αH2B, αH3, and αH4) to histones (H2A, H2B, H3, and

H4), respectively. Figure 30 shows αH2A, αH2B, αH3, and αH4 produced a strong

supershift with both canonical and the HMGB1-remodeled nucleosomes (N’/N”), clearly

indicating the presence of all (H2A, H2B, H3, and H4) core histones.

The anti-histone antibodies (H2B, H3, and H4) were purchased from Active Motif.

Histone H2B (Cat # 39125) has a molecular weight of 15 kDa. The epitope is peptide

including the C-terminal region of histone H2B. Histone H3 (Cat # 39163), molecular

weight 17 kDa; the epitope was C-terminal peptide of histone H3. Histone H4 (Cat #

39269) molecular weight 8 kDa; epitope was human histone H4. Histone H2A (Cat # 07-

146) was purchased from Millipore, molecular weight was 14 kDa. The epitope is KLH-

conjugated synthetic peptide corresponding to amino acids 88-97 of human histone H2A.

Figure 30. Supershift assay to determine the presence of core histones in nucleosomes, and HMGB1-remodeled nucleosomes: Lane 1, nucleosomes in TE/sucrose buffer; Lanes

2-5, nucleosomes in TE/sucrose buffer reacted with -H2A, -H2B, -H3, and -H4, respectively. Lane 6 contains N’/N” in TE/sucrose buffer; lane 7-10, remodeled nucleosomes in TE/sucrose buffer with -H2A, -H2B, -H3, and -H4, respectively.

Nucleosomes were incubated on ice with 2 L of anti-histone antibody (-H2A, -H2B, -

H3, and -H4) for 10 minutes in10µL reaction volume. After the incubation, they were loaded onto an EMSA gel and run for 2 hours at 200V at 4oC.

13. ER binding to HMGB1-remodeled nucleosomes (N’/N”)

The remodeled nucleosome population was used in binding studies with ER. Figure

31A shows that the remodeled structures were able to bind to ER without the addition of

HMGB1. The N’/N” band disappeared as the ER concentration was increased and formed an ER complex with N’/N”. This shows that the action of HMGB1 was able to change the character of the nucleosomes, to permit ER binding. The KD was less than 20 nM.

Interestingly, figure 31B shows that the presence of 1 ng/ µL poly (dI-dC) in the ER binding buffer caused both N’/N” to revert to the same mobility as the canonical nucleosome. Both N’/N” forms are, therefore, unstable in the presence of this polyelectrolyte environment. The KD in the presence of poly (dI-dC) was higher, with the

KD of 40 nM indicating a weaker binding of ER. Figure 32 shows the binding profile from three separate experiments, of ER binding to remodeled nucleosomes. The graph was plotted dividing the amount of complex formed by the total amount of DNA (See chapter

2).

Figure 31. ER binding to remodeled nucleosomes in the absence (A) and presence (B) of 1 ng/ µL poly (dI-dC). Nucleosomes were incubated with increasing concentration of

ER in ER binding buffer (table 5). The reaction was incubated on ice for 30 minutes and run for 2 hour on 4% polyacrylamide gel. Lane 1, nucleosomes alone; lane 2-9, remodeled nucleosomes binding to ER concentrations of 0, 10, 20, 30, 40, 50, 60 and 80 nM, respectively. Each lane in figure B contained 1 ng/ µL of poly (dI-dC). 93

100

% Complex 20

110100 ER (nM)

Figure 32. Binding profile of remodeled nucleosomes in the absence (■) and presence

(●) of poly (dI-dC). The Kd values were plotted from three independent gels, using the

Origin 6.2 software. ER concentrations were 0, 10, 20, 30, 40, 50, 60 and 80 nM.

14. DNase I digestion of nucleosomes and N’/N”

To further characterize the HMGB1-remodeled nucleosomes (N’/N”), DNase I

digestion was performed on both canonical and N’/N” nucleosomes. Each of the

nucleosomal DNAs, 2E2, 3E1 and 4E0 used in the experiment were labeled at the EcoRI

end. We have already showed that HMGB1 changes the character of the nucleosome to

allow for the binding of ER. Therefore, we wanted to determine if the rotational phasing

within the nucleosome, as reflected in the altered DNase I 10 bp patterns was evident.

Figures 33, 34 and 35 show that the basic DNase I 10 bp ladder is maintained on

both the canonical and each of N’/N”. Therefore, the rotational phasing of the nucleosome

remains unchanged in the remodeled nucleosome. However, figures 33, 34 and 35 show

that the remodeled nucleosomes exhibit additional bands on the gel compared to the

canonical nucleosomes. Those sites on the N’/N” have increased sensitivity to DNase I

cutting which are not observed in the canonical nucleosomes nor in the nucleosomes exposed to 400 nM HMGB1. This suggests that the structures of N’/N” are different from the nucleosomes, and the minor grooves are more accessible at those sites that show increased DNase I sensitivity

Figure 33 (DNase I on 2E2) shows five additional bands in the remodeled nucleosomes which were not present in the canonical nucleosomes. The locations of the bands were 56, 74, 92, 97 and 109 bp.

Figure 34 (DNase I on 3E1) shows seven additional bands in the remodeled nucleosomes which were not present in the canonical nucleosomes. The locations of the bands ranged between 70 – 130 bp (70, 90, 92,120, 125, 132 and 140 bp). 95

Figure 35 (DNase I on 4E0) showed ten additional bands in the N’/N” and two hypersensitive sites. The locations of the bands were 50, 51, 52, 61, 62, 71, 79, 93, 94 and

102 bp.

There was however, no evidence of additional bands observed at the position of the cERE. The location of the additional bands for each nucleosomal DNA is presented in

Table 6. Figure 36 shows a comparison of the extra DNase I sensitive sites for the 3 nucleosomal DNAs. Most of the additional DNase I sensitive sites are centered around mid portions of the DNA (50-110), with a few cuts showing up at the end of the DNA. Most reside in the NPSs, except 3E1. This suggests that even though specific cuts were not observed in the cERE sites, the immediate vicinity of the cERE becomes more accessible when the nucleosomes are remodeled by the HMGB1. This may explain why ER is able to bind to nucleosomes in the presence of HMGB1. 96

Figure 33. DNase I digestion of canonical nucleosomes and N’/N” (2E2). Lane 1: A/G ladder, lanes 2-4: nucleosomes and lanes 5-7: N’/N” digested with 0.04U DNase I for 30,

60 and 120 seconds, respectively. The reaction was carried out at 4 oC to maintain the integrity of the N’/N”. The arrows on the right of the picture (←) indicate additional DNase

I cuts on the modified nucleosomes, cutting sites on the modified nucleosomes that are different from the canonical nucleosomes. The arrows on the left (→) indicate the DNase I

10 bp cutting sites on nucleosomes. The bar on the left represents the cERE (74-88 bp). 97

Figure 34. DNase I digestion of canonical nucleosomes and N’/N” (3E1). Lane 1: A/G ladder, lanes 2-5: nucleosomes and lanes 6-9: N’/N”; digested with 0.04U DNase I for 30,

60, 90 and 120 seconds, respectively. The reaction was carried out at 4 oC to maintain the integrity of the N’/N”. The arrows on the right of the picture (←) indicate additional DNase

I cuts on the modified nucleosomes, cutting sites on the modified nucleosomes that are different from the canonical nucleosomes. The arrows on the left (→) indicate the DNase I

10 bp cutting sites on nucleosomes. The bar on the left represents the cERE (94-108 bp). 98

Figure 35. DNase I digestion of canonical nucleosomes and N’/N” (4E0). Lane 1, A/G

ladder; lanes 2-5, nucleosomes and lanes 6-9, N’/N”; digested with 0.04U DNase I for 30,

60, 90 and 120 seconds, respectively. The reaction was carried out at 4 oC to maintain the

integrity of the N’/N”. The arrows on the right of the picture (←) indicate additional DNase

I cuts on the modified nucleosomes, cutting sites on the modified nucleosomes that are

different from the canonical nucleosomes. The arrows on the left (→) indicate the DNase I

10 bp cutting sites on nucleosomes. The bar on the left represents the cERE (114-128 bp).

The dots on the right are the hypersensitive DNase I cutting sites. 99

Location of hypersensitive and new bands (bps) DNA in N’/N”

2E2 56, 74, 92, 97 and 109

3E1 70, 90, 92,120, 125, 132 and 140

4E0 50, 51, 52, 61, 62, 63, 71, 79, 93, 94, 102 and 103

Table 6. Positions of additional DNase I cuts in DNA of N’/N” nucleosomes. The table shows the appearance of 5 bands for 2E2, 7 bands for 3E1 and 10 additional bands and 2 hypersensitive sites (bold numbers) for 4E0 N’/N”. 100

Figure 36. Comparison of DNase I sensitive sites on nucleosomal DNAs. The scheme shows the location of additional DNA bands in N’/N” from DNase I gel. D refers to dyad axis. There were 5 bands (56, 74, 92, 97 and 109 bp) for 2E2, 7 bands (70, 90, 92,120,

125, 132 and 140 bp) for 3E1 and 12 additional bands (50, 51, 52, 61, 62, 63, 71, 79, 93,

94, 102 and 103 bp) for 4E0 modified nucleosomes (63 and 103 are hypersensitive sites).

The dark horizontal band represents the position of the cERE. The dashed box represents

60 bp of DNA interaction with histone H3/H4 tetramer in the nucleosome. Thirty base pairs to either side of the dashed box interacts with H2A/H2B while the last 13 bp of DNA at the entry and exit point of the nucleosome is organized exclusive by the α-helical histone fold extension of H3 and the preceding H3 N-terminal tail (Luger and Richmond, 1998). Most additional DNase I sensitive cuts occur within the NPSs and none within the cERE.

101

15. HMGB1 does not alter the translational position of DNA within nucleosomes

We asked whether HMGB1 would change the translational position of DNA within the nucleosome, and therefore, allow ER to bind to cERE.

Exonuclease III cuts a single strand of double stranded DNA from the end in the 3’ to 5’ direction. The enzyme pauses when it encounters a protein or nucleosomal proteins. If

HMGB1 changes the translational position of DNA within the nucleosome, we would expect to find that Exo III digests the nucleosomal DNA into shorter fragments compared to the (HMGB1) untreated nucleosomes. Exonuclease III digestion was therefore, performed at room temperature after nucleosomes were incubated with 1600 nM HMGB1 on ice for 1 hour. Purified DNA was run on 8% denaturing gel for 3 hours.

Exo III reaction with 4E0 DNA produced cleavage and a series of shorter DNA fragments in the absence of HMGB1 (figure 37, lane 2) compared to the pattern in the presence of 1600 nM HMGB1 (figure 37, lane 3). This indicates that the Exo III extensively cut DNA in the absence of HMGB1. HMGB1 at a concentration of 1600 nM appears to significantly inhibit the activity of the Exo III enzyme (figure 37). However,

Exo III on HMGB1-treated nucleosomes gave virtually the same profile for nucleosomes as untreated nucleosomes with HMGB1, suggesting that HMGB1 had no effect on the translation of DNA. A change in the translational position of DNA in the nucleosomes would have resulted in the exonuclease gaining greater access to the DNA in the presence of 1600 nM HMGB1. Therefore, the HMGB1 does not appear to alter the translational position of DNA on the nucleosomes. However, the inhibitory effect of the HMGB1 on

Exo III activity limits the certainty of this conclusion.

102

Figure 37. Exo III digestion on nucleosomes (4E0), with and without 1600 nM

HMGB1. Nucleosomes were incubated with or without 1600 nM HMGB1 on ice for 1

hour. The reaction was transferred to room temperature for 5 minutes and digested with

1U/µL of Exo III. The reaction was stopped at 1, 2, 4 and 8 minutes. The DNA fragments

were purified with phenol chloroform and ethanol precipitated. The radioactivity was normalized for each well and the DNA dissolved in 10% formamide. An 8% denaturing gel was run for 3 hours at 1500V. Lane 1, 161 bp DNA; lane 2, DNA; lane 3, DNA with 1600

nM HMGB1, lane 4-7, nucleosomes; lanes 8-11, nucleosomes with 1600 nM HMGB1. 103

16. Ava I restriction enzymes digest of nucleosomes and modified nucleosomes

Another test of the character and accessibility of the nucleosomal DNA is to investigate the accessibility of a restriction enzyme to its recognition site located at different regions within the nucleosomal DNA. To determine if increases the accessibility of DNA in nucleosomes, accessibility of Ava I to its recognition site was determined for the canonical and HMGB1-modified nucleosomes. There is one Ava I recognition sequence present in each DNA construct (Ava I site; 2E2 is 98 bp, 3E1 is 118 bp and 4E0 is 138 bp from the EcoRI end of the labeled DNA).

A measure of Ava I accessibility of the site was determined by measuring the rate of cleavage and the extent to which Ava I can cleave the DNA within the nucleosome. Figure

38 shows the schematic of position of cERE and the AvaI restriction recognition sequence cutting sites (C/TCGGG). Table 7 indicates the size of the radiolabeled DNA produced by

Ava I cutting for the three DNA constructs. Figure 39A shows the gel for the digestion as a function of time, while figure 39B show a plot of the rate and extent of digestion. Figure

39B shows that after 5 min of digestion, 24% of DNA was cleaved in the unmodified nucleosomes, while 60% of the N’/N” was cleaved. After 30 minutes, 41% of total unmodified nucleosomal DNA was cut compared to 75% in the remodeled nucleosome.

This may be explained by the addition of 1600 nM HMGB1 to the nucleosomes results in remodeling of the nucleosome to change the DNA-histone interactions, making the new population of nucleosomal DNA more accessible to Ava I at 138 bp (or 23 bp) from the

DNA end. Figure 39B showed the rate and extent of digestion (from phosphoimager intensities) which was calculated from the amount of DNA recovered in the experiment as a function of total DNA. 104

Figure 38. Schematic of DNA showing the general position of cERE and the Ava I

restriction recognition sequence and cutting sites in all DNAs.

DNA Size of labeled DNA after Ava I Center of cERE from cleavage labeled end (bp) 2E2 98 83

3E1 118 103

4E0 138 121 Table 7. The size of radiolabeled DNA after Ava I digestion and the center of cERE from the labeled end of DNA.

105

Figure 39. Ava I digestion of 4E0 nucleosomes and N’/N”. Each of the nucleosomal

DNAs in the experiment was labeled at the EcoRI end. Equal counts (cpm) of nucleosomal

DNA in nucleosomes and remodeled nucleosome were incubated in a 200 µL total volume, and digested with 100 U of enzymes. Aliquots of 40 µL were taken at 0, 5, 10, 15 and 30 minutes and EDTA added to a final concentration of 25 mM. (A) The purified labeled 106

DNA from the digested nucleosomes was run on 8% denaturing gel for 90 minutes. (B)

Kinetic plot of the percent Ava I digestion versus time for nucleosome and N’/N” forms.

The plot shows the time profile for the rate and extent of digestion of DNA that was cut by

Ava I to produce the DNA fragment of 138 bp in the canonical (♦) and modified

nucleosomes (■). The graph represents values of two separate experiments.

17. Effect of temperature on HMGB1- remodeled nucleosomes (4E0)

The thermal stability of the canonical and remodeled nucleosomes was determined by incubating the nucleosomes at three different temperatures. Aliquots of remodeled nucleosomes were incubated at 4 oC, 37 oC and 55 oC, and compared with canonical

nucleosomes incubated at 37 oC. Figure 40 shows that the canonical nucleosomes were

stable at 37 oC, even if the incubation time was prolonged to overnight. The N’/N” were

stable at 4 oC at all times examined from 0-120 minutes and overnight. Our experience with

this is that, the N’/N” are stable for at least a month at -20 oC. However, the N’/N” were

unstable at 37 oC and 55 oC, as the band for N” reverted back to N’. Therefore, the N’

appears to be more thermodynamically stable than N”. There is, however, no evidence to

show N’ reverting any further to canonical nucleosomes. Therefore, the N’ form is stable

for an extended time at 37 oC and 55 oC. 107

Figure 40. Effect of temperature on mobility of nucleosomes and remodeled (4E0) nucleosomes. Lanes 1-4, nucleosome in TE/sucrose buffer at 37 oC for 0, 60, 120 minutes and overnight (O/N); lanes 5-8, remodeled nucleosomes in TE/sucrose buffer at 4 oC for 0,

60, 120 minutes and overnight (O/N); lanes 9-12, remodeled nucleosomes in TE/sucrose buffer at 37 oC for 0, 60, 120 minutes and overnight (O/N); lanes 13-16, remodeled nucleosomes in TE/sucrose buffer at 55 oC for 0, 60, 120 minutes and overnight (O/N).

After the incubation, a 10 µL aliquot of each of nucleosomes or remodeled nucleosomes was loaded onto an EMSA gel and run for 2 hours at 200V at 4 oC.

108

18. Effect of increasing salt concentration on HMGB1- remodeled nucleosomes

The stability of the modified nucleosomes in increasing NaCl concentration was determined by incubating modified nucleosomes with 0-300 mM NaCl. Unlike heat, salt converts both N’ and N” back to the canonical nucleosomes at 100 mM NaCl (figure 41).

The result indicates that the remodeled nucleosomes became increasingly unstable in NaCl and was completely reverted to canonical nucleosomes at 100 mM NaCl. 109

Figure 41. Effect of increasing NaCl concentration on mobility of remodeled nucleosomes. Lane 1, DNA; Lane 2, nucleosome in TE/sucrose buffer; Lane 3, remodeled nucleosome in TE/sucrose buffer; Lanes 4-8, remodeled nucleosomes in TE/sucrose buffer with mM NaCl as indicated. D (DNA), N (nucleosomes) and N’/ N” (HMGB1-remodeled nucleosomes). After incubation on ice for 15 minutes, the salt concentrations in the reaction mixture were diluted to 100 mM for solutions in lanes 7 and 8. Those in lanes 4 and 5 (25 and 50 nM) were kept at the same NaCl concentration. After incubation, 20 µL aliquots of each sample were loaded onto an EMSA gel and run for 2 hours at 200V at 4 oC.

110

19. Stability of HMGB1-remodeled nucleosomes in different buffers

Previous experiment showed that the HMGB1-remodeled nucleosomes were unstable in high salt environment (see section 18). The buffers for Exo III digestion and ER binding reactions represent a high salt environment, and therefore might affect the stability of remodeled nucleosomes. To determine the stability of HMGB1-remodeled nucleosomes in different components of ER dilution buffer, the HMGB1-remodeled nucleosomes were incubated on ice with different components of the ER buffer, DNase I buffer, Exo III buffer, and Exo III buffer (Fisher Scientific), for 30 minutes and the reaction mixture was run on 4% polyacrylamide gel. Ten microliter aliquots of nucleosomes or HMGB1- remodeled nucleosomes (N’/N’’) were treated with 10 L of each of the buffers. DNase I buffer (10 mM Tris-HCl, pH 7.5, 2.5 mM MgCl2, 0.5 mM CaCl2), Exo III buffer (33 mM

Tris- acetate, pH 7.8, 66 mM KAc, 10 mM MgAc2, 0.5 mM DTT), Exo III buffer from

Fisher Scientific (66 mM Tris-HCl, pH 8, 0.6 mM MgCl2), TE/sucrose (18%) buffer (10

mM Tris-HCl, pH 8.0, 1 mM EDTA, 18% sucrose) and ER dilution buffer (80 mM KCl,

10% glycerol, 15 mM Tris-HCl, pH 8.0, 0.2 mM EDTA, 0.4 mM DTT, 100 ng/µL BSA, 2 ng/µL poly (dI-dC). The final reaction buffer concentration for N’/N” in ER dilution buffer is presented in chapter 2.

Figure 42 shows that the canonical nucleosomes are very stable and do not change mobility or dissociate into DNA and histones in this high ionic environment. However, although the N’/N” is stable in TE/sucrose (lane 6) and DNase I buffered (lane 8), it is unstable in ER dilution buffer (lane 7) and Exo III buffer (lanes 9), and the N’/N” form reverts to canonical nucleosomes. There is however, no evidence of dissociation into free

DNA and histones. This suggests that the N’/N” are very sensitive to buffer conditions. 111

Table 5. Final concentration of each component of the reaction buffer

DNase I Tris-HCl 10 mM

100 mM Tris-HCl (pH 7.5), 20 mM MgCl2 MgCl2 4 mM and 5 mM CaCl 2 CaCl 0.5 mM 2 Exo III** Tris-acetate 3.3 mM

MgAc 1.0 mM 33 mM Tris- acetate, pH 7.8, 66 mM KAc, 2 KAc 6.6 mM 10 mM MgAc , 0.5 mM DTT 2 2 DTT 0.05 mM *Note: The reaction mixture was prepared by mixing one part ER dilution buffer and one part nucleosomes in 18% sucrose/TE buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA). Therefore, the final concentration of sucrose is 9% and glycerol is 5% in all the reactions **Exo III buffer converts N’/N” to canonical nucleosomes and was therefore, not used in Exo III digestion. All Exo III digestions were done in DNase I buffer which kept the Exo III activity and did not convert any of N’/N” to canonical form. 112

Figure 42. Stability of HMGB1-remodeled nucleosomes in different buffers. Lanes 1-

5, nucleosomes in TE/sucrose buffer, ER buffer, DNase I buffer, Exo III buffer, Exo III buffer (Fisher Scientific), respectively; lane 6, remodeled nucleosomes in TE/sucrose buffer; lane 7, ER buffer; lane 8, DNase I buffer; lane 9, Exo III buffer; lane 10, Exo III buffer (Fisher Scientific). The reaction was incubated on ice for 30 minutes and run for 2 hours on 4% polyacrylamide gel. 113

20. Effect of increasing levels of unlabeled DNA on EMSA mobility of modified nucleosomes

We asked how stable the modified nucleosomes were in the presence of excess

DNA. This unlabeled 161 bp DNA was the same DNA used to prepare the nucleosomes. It has been shown that incubation of nucleosomes with excess DNA can disrupt nucleosomes and allow for the formation of free labeled DNA, dissociated from the nucleosomes

(Workman and Kingston, 1992; Juan et al, 1993). Bands corresponding to free labeled

DNA may be expected if excess cold DNA competed for the core histones and facilitated

DNA dissociation from the nucleosomes. However, figure 43 showed that there was no evidence of DNA dissociating from the histones. The remodeled nucleosomes were unstable in the excess cold DNA, and reverted back to the canonical nucleosomes. This is similar to our finding with the presence of poly (dI-dC), but with the 161 bp DNA, the concentration can be determined. 114

Figure 43. Effect of increasing the levels of unlabeled 161 bp DNA on modified nucleosome. The modified nucleosomes were incubated with increasing amounts of unlabeled DNA on ice for 30 minutes and run for 2 hours on 4% polyacrylamide gel. Lane

1, DNA; lane 2, nucleosomes; lane 3, remodeled nucleosome from sucrose gradient; lane 4-

8, remodeled nucleosome treated with unlabeled DNA (nM) as indicated. D (DNA), N

(nucleosome) and N’ and N” (HMGB1-remodeled nucleosome).

115

21. Determine if the continuous presence of HMGB1 is required to maintain the

stability of remodeled nucleosomes

We have previously shown that N” and N’ are unstable in a high salt environment.

At a salt concentration of 50 nM, both complexes revert to canonical nucleosomes (figure

41). We have also shown previously that 1 µg of anti-HMGB1 which is equivalent to 6.6 *

10 -12 moles in 10 µL is sufficient to bind to 400 nM HMGB1. In the remodeling reaction, nucleosomes were treated with 1600 nM and separated on a 5 mL sucrose gradient. If one could assume that during the sedimentation of the HMGB1-remodeled nucleosomes, the

HMGB1 was evenly distributed in the sucrose gradient, then the greatest HMGB1 concentration through out the gradient would be 100 nM (1.0 * 10-7 moles/L). In this case,

we would expect that in a reaction with 100 nM HMGB1 (10µL), 1 µg of anti-HMGB1 in

10 µL reaction volume would bind and eliminate all the HMGB1 in a reaction. If HMGB1

is essential for holding the remodeled nucleosome together, addition of anti-HMGB1

would bind to the HMGB1 and eliminate those molecules from the reaction. This would

produce an EMSA band shift of the nucleosome back to the position of the canonical nucleosome.

Figure 44 shows that the addition of 1µg anti-HMGB1 to the HMGB1-remodeled nucleosomes induced an insignificant shift of the N’/N” complex (fig 44, lane 2 and 3).

This suggests that there was not enough HMGB1 present to begin with to affect the N’/N”.

It also suggest that any small level of HMGB1 that may be present with the remodeled

nucleosomes is not sufficient to keep the N’/N” complex intact. However, N’/N” complex

revert canonical nucleosome at high concentration (4µg) of antibody to HMGB1 (data not

shown). This is, however, due to the high salt environment (40 mM final concentration) 116 and not the antibody since antibody buffer lacking antibody produced the same effect. The antibody was supplied in 0.1 M Tris-glycine, pH 7.4, 0.15 M NaCl, 0.05% sodium azide and 30% glycerol. 117

Figure 44. Titration of HMGB1-remodeled nucleosome with anti-HMGB1. HMGB1- remodeled nucleosomes were incubated on ice for 30 minutes with 1µg of α HMGB1 or α

HMGB1 buffer (heat the αHMGB1 at 78 oC for 20 minutes to denature the protein and spin it down). Lane 1, canonical nucleosome; lane 2 and 3, contained 0 and 1µg of αHMGB1; lanes 4 and 5 contained the same amount of α HMGB1 buffer as lanes 2and 3, so we can eliminate any effect that may be due to the buffer. 118

22. Characteristics and reactions with tailless nucleosomes

The histone tail domains are the regions of the protein that resides outside the core

of the nucleosome. These are located at the N-terminal of all the core histones; in addition

to the C-terminal of histone H2A. Removal of the tails does not affect the stability of translational or rotational position of the nucleosome (Dong et al, 1989, Hayes et al, 1991).

To explore if the tail domains influence the action of HMGB1, chromatin digested with micrococcal nuclease was isolated and then the oligonucleosomes were digested with trypsin to produce histones in which the tails were cleaved off (See chapter 2) (Mutskov et al, 1998). Figure 45 shows the SDS-PAGE of the tailless histones in the tailless nucleosome with molecular weight less that that of H4 (Hayes et al, 1991). The tailless oligomers were then used in the exchange reaction to produce tailless nucleosomes which

was then purified by 5-30% sucrose gradient sedimentation as previously described for

canonical nucleosomes.

22.1 Characteristics of tailless nucleosomes

The tailless nucleosomes were reconstituted and isolated as before on 5-30%

TE/sucrose buffer. Figure 47 shows that tubes 10-12 contained the peak of radioactivity

and were the same fraction as those nucleosome fractions collected in the tailed

nucleosomes. Therefore, tailless nucleosomes sediment at the same density as those in

tailed nucleosomes. EMSA indicates that the tailed nucleosome have a slightly greater

mobility compared to the tailless nucleosome (data not shown). This is an unexpected

result since the tailless histones are smaller in size, contain reduced number of positive

charges, and therefore, it is expected to move faster in a gel (keeping current constant). 119

Figure 47 shows that the tailless nucleosomes were separated from the unincorporated

DNA on 5-30% TE/sucrose gradient (M and M). Aliquots of 250 µL were collected and stored at –20 oC. 120

Figure 45. SDS-PAGE of core and tailless histones in oligonucleosomes. Lanes 1, molecular weight markers; lane 2, trypsin digested chromatin extract (tailless histones); lane 3, core histone; lane 4, fractionated oligonucleosomes (with H1). The results show low mobility bands about 10 kDa in size representing the tailless histones. Only four bands representing the core histone proteins (H2A, 13.9 kDa; H2B, 13.7 kDa; H3, 15 kDa and

H4, 11 kDa). The bands for histones run slightly different from the molecular weight markers because histone proteins are highly charged. H1 with molecular weight of 21 kDa appears at ca. 27 kDa in lane 4. 121

3500

3000 2500

m 2000

1500

Relative cp Relative 1000

500

0 0 2 4 6 8 10 12 14 16 18 20 Fraction Number

Figure 46. Sucrose gradient sedimentation profile showing free DNA and tailless

nucleosomes. Individual fractions were collected from 5-30% linear sucrose gradient by

pushing a 50 % sucrose solution from the bottom of the 5 mL centrifuge tube. Samples

were collected (250 µL) and the counts per minute determined with the DuPont BC 2000

counter. Fraction 5 contains labeled DNA, while fractions 10-12 contain tailless

nucleosomal DNA. The fractions were stored at -20 oC. 122

Figure 47. Sucrose gradient centrifugation fractions analyzed on polyacrylamide gel.

About a 10 µL sample of the various sucrose gradient fractions was run on a 4%

polyacrylamide gel. The bands corresponding to tailless nucleosomes were not contaminated with free DNA.

123

22.2 Effect of removing histone tails from core histones

We hypothesized that the histone tails may be involved in the HMGB1 effect on

accessibility of the DNA. This may occur by the positively charged tails binding to DNA

and interfering with ER/ERE binding or restricting Ava I binding access to or cutting the

nucleosomal DNA. To test this hypothesis, we first looked at the effect of removal of the

histone tails on ER binding affinity.

Figure 48A shows that ER binds with a significantly higher binding affinity to the

tailless nucleosomes compared with tailed nucleosome. The KD for ER binding to tailless

nucleosome without HMGB1 was 45 nM compared to an estimated KD ~ 300 nM for

normal nucleosomes. The addition of 400 nM HMGB1 further enhanced ER binding to tailless nucleosomes, with a KD of 25 nM compared to a KD value of 50 nM for canonical

nucleosome having intact core histone tails (figure 48B). Therefore, removal of the tails

from the core histones clearly enhanced the binding of ER to nucleosomes. The values of

three separate experiments were used to plot the binding profile of ER binding to tailless

nucleosomes in figure 49. The graph was plotted dividing the amount of complex formed by the total amount of DNA (See chapter 2). 124

Figure 48. ER binding affinity to tailless nucleosomes (4E0) in the (A) absence of

HMGB1 and in the (B) presence of 400 nM HMGB1. Nucleosomes were incubated with increasing ER concentrations on ice for 30 minutes and run on 4% polyacrylamide gel for 2 hours. Lanes 1-8 contained 0, 10, 20, 30, 40, 50, 60 and 80 nM ER, respectively. 125

100

% Complex 20

0 110

ER (nM)

Figure 49. Binding profile of ER binding to tailless nucleosomes in the absence and in

the presence of 400 nM HMGB1. The dots (●) indicate binding in the presence of 400 nM and squares (■) indicate binding of ER in the absence of HMGB1. 126

22.3 DNase I footprint of ER/cERE in 2E2 tailless nucleosomes

A DNase I footprint was performed on the binding of ER to tailless nucleosomes in the presence or absence of 400 nM HMGB1 to determined whether the binding of ER was sequence specific. Figure 50 shows a cERE footprint when 100 nM ER was reacted with

2E2 nucleosomal DNA, in the absence or presence of 400 nM HMGB1. Figure 50 shows that there is a decrease in cutting at 74-88 bp, which is the location of the cERE, from the

EcoRI end. Therefore, this indicates that ER binds to cERE and reduces the accessibility of the site to DNase I cutting. HMGB1 further enhances the binding of ER to the tailless nucleosome. This is evident by comparing lanes 3 and 4 to 6and 7, figure 50. The footprint for ER binding to the cERE is apparent at the low ER concentration of 50 nM in the presence of 400 nM HMGB1 as compared to 100 nM in the absence of HMGB1. 127

Figure 50. DNase I footprint of tailless nucleosomes (2E2) in the absence and presence

of 400 nM HMGB1. The tailless nucleosomes were incubated on ice for 30 minutes with

or without 400 nM HMGB1 and increasing concentration of ER. The reactions were then

kept at room temperature for 5 minutes and digested with 0.04U DNase I. The reaction was stopped after a minute of digestion with DNase I by increasing the EDTA concentration to 25 mM. The purified DNA was run on 8% polyacrylamide gel for 2.5 hours. Lane 1, AG ladder; lane 2-4, tailless nucleosomes binding to 0, 50 and 100 nM ER in the absence of 400 nM HMGB1; lanes 5-7, tailless nucleosomes binding to 0, 50 and

100 nM ER in the presence of 400 nM HMGB1. The bar represents the location of the

cERE (74-88 bp). 128

23. Effect of increasing HMGB1 concentration on tailless nucleosome

To determine if increasing levels of HMGB1 had increasingly altered nucleosome

mobility as determined in normal nucleosomes, the tailless nucleosomes were incubated

with 400, 800 and 1600 nM HMGB1 for one hour at 4 oC. The reaction mixtures were then

analyzed on 4% polyacrylamide gel to determine if HMGB1 changed the EMSA mobility of the tailless nucleosomes. Figure 51 shows that mobility of nucleosome bands increasingly decreased with increasing levels of HMGB1 in the reaction, as also observed

for canonical tailed nucleosomes.

Previous work showed that HMGB1 did not form a stable component of the newly

formed complex, as reflected in no supershift of the bands on addition of anti-HMGB1

(Sarpong, 2006). Therefore, the reduced mobility of the newly formed complex in the

presence of HMGB1 could not be due to HMGB1 binding to the nucleosomes, and

changing its mobility. The highly positively charged tails may play a role by transiently

interacting with the acidic tail of the HMGB1 and the negatively charged DNA interacting

with the Aand B boxes of the HMGB1. 129

Figure 51. EMSA mobility of tailless nucleosomes with increasing levels of HMGB1.

The tailless nucleosomes were incubated with 0, 400, 800 and 1600 nM HMGB1 on ice for an hour and run on 4% polyacrylamide for 2 hours.

130

24. Isolation of HMGB1-remodeled tailless nucleosomes (N*)

To further characterize the lower mobility band formed in the presence of 1600 nM

HMGB1, an attempt was made to isolate the complex from a sucrose gradient. Tailless nucleosomes were treated with 1600 nM HMGB1 on ice for an hour and the contents of the tube were loaded onto a 5-30% sucrose gradient. Fractions were collected as before, and analyzed on 4% polyacrylamide gel. We have previously shown that two distinct populations of HMGB1 remodeled nucleosomes (N’ and N”) were produced in the case of normal nucleosome (figure 29). We observed a difference in the tailless nucleosomes compared to the normal nucleosomes. Figure 52 shows a single unresolved (broad)

HMGB1 remodeled tailless nucleosome band, compared to two distinct resolved bands for the normal remodeled nucleosomes (N’/N”). These data suggest that the tails may be involved in the formation of the two distinct populations (N’/N”) formed in the canonical nucleosomes. The highly positively charged tails may play a role by transiently interacting with the acidic tail of the HMGB1 and the negatively charged DNA interacting with the

Aand B of the HMGB1. Without the tails, the interaction may be between the DNA and

HMGB1 alone. Fractions from lanes 21 and 22 were used for subsequent reactions. The N* are stable for a month when stored in TE/sucrose buffer at -20 oC.

131

Figure 52. Sedimentation profiles of HMGB1-remodeled tailless nucleosomes (4E0).

Tailless nucleosomes were treated, with 1600 nM HMGB1 and sedimented on 5-30%

sucrose gradient for 16 hours. Aliquots from each fraction were loaded on 4% gel and

electrophoresed for 2 hours at 200V. Nt is tailless nucleosome and N* is remodeled tailless nucleosome. 132

25. The influence of poly (dI-dC) on ER binding to N*

The HMGB1 remodeled tailless nucleosomes were isolated (fractions 21 and 22)

and used in binding studies with ER. Figure 53A showed that the remodeled nucleosomes were able to bind to ER without the addition of addition HMGB1. The N* band disappeared as the ER concentration was increased and formed ER complex with HR-tNS.

Figure 41 A shows that HMGB1 was able to change the character of the nucleosomes, to facilitate stronger ER binding to cERE. Figure 53B showed that the addition of 1 ng/ µL poly (dI-dC) in the ER binding buffer forced the N* to revert to the same mobility as the tailless nucleosome. This is an effect observed similarly with remodeled N’/N” with tails.

The N* is therefore, unstable in a high electrolyte environment. The KD was less than 20

nM in the absence of poly (dI-dC) and about 30 nM in the presence of poly (dI-dC). Figure

54 shows the Kd values plotted from three independent gels, using the Origin 6.2 software. 133

Figure 53. ER binding to remodeled tailless nucleosomes in (A) the absence and (B) presence of poly (dI-dC) (4E0). The reaction was incubated on ice for 30 minutes and run for 2 hours on 4% polyacrylamide gel. Lane 1, tailless nucleosomes; lane 2, remodeled tailless nucleosomes; lanes 3-9, remodeled tailless nucleosomes binding to ER concentrations of 10, 20, 30, 40, 50, 60 and 80 nM respectively. Each lane in figure B contained 1 ng/ µL of poly (dI-dC). 134

100

% Complex 20

110100 ER (nM)

Figure 54. Binding profile of remodeled nucleosomes in the absence (■) and presence

(●) of poly (dI-dC (4E0). The Kd values were plotted from three independent gels, using the Origin 6.2 software. ER concentrations were 0, 10, 20, 30, 40, 50, 60 and 80 nM.

135

26. DNase I digestion of tailless nucleosomes and N*

We have already shown that HMGB1 1) changes the character of nucleosomes as

shown by reducing its gel mobility and 2) enhances the binding affinity of ER. To further

characterize the remodeled structures in the presence of HMGB1, DNase I digestion was

performed on tailless and N* nucleosomes. We anticipated that the DNase I pattern may

show a difference between the tailless nucleosomes and the remodeled tailless nucleosomes.

Figure 55 and 56 shows that the 10 bp ladder is maintained even after the cleavage

of the tails, consistent with the notion that the tail domains do not influence the rotational phasing of the nucleosome (Lee et al, 1993). Therefore, the rotation of the nucleosomes does not change with the removal of the histone tails for the core histones. The pattern of

DNase I 10 bp cleavage pattern is the same for both tailless and N*, with additional bands appearing in the remodeled nucleosomes. Figure 55 (tailless 2E2) shows twelve additional bands in the N* which were not present in the tailless nucleosomes. Two of the additional cuts appear in the position of the cERE (82 and 83 bp). The locations of the bands are at 50,

51, 52, 59, 60, 61, 62, 71, 82, 83, 92 and 98 bp. Figure 56 (tailless 4E0) also shows twelve additional bands in the HR-tNS which were not present in the tailless nucleosomes. The locations of the bands are at 50, 51, 52, 59, 60, 61, 82, 83, 84, 104, 112 and 140 bp. This again suggests that the structure of HR-tNS is very different from the normal tailless nucleosomes and that the minor grooves are more accessible at those locations which show

DNase I sensitivity. There was however, no evidence of additional bands observed in the position of the cERE. 136

Table 8 shows a comparison of the location of the additional bands in the tailless nucleosomes with that normal nucleosome. DNase I cut sites in both tailless and tailed nucleosomes (remodeled) were observed from 50-140 bp of the DNA. This may explain why ER is able to bind more easily to the remodeled nucleosomes compared to canonical nucleosomes. Figure 57 shows a layout of DNase I sensitive sites on linear DNA of all the three DNA constructs compared to HR-tNS. The N* shows DNase I sensitive sites opened along the entire length of the DNA. The additional bands observed in N* may help rationalize why N* is the most accessible to the Ava I digestion (figure 56) and also binds strongly to ER. 137

Figure 55. DNase I of tailless nucleosomes and N* (2E2). Lane 1, A/G ladder; lanes 2-4, nucleosomes digested for 30, 60 and 90 seconds, respectively; lanes 5-7, modified nucleosomes digested for 30, 60 and 90 seconds, respectively. The reaction was performed at 4 oC to maintain the integrity of the modified nucleosomes. The arrows on the right of the picture (←) indicate additional DNase I cut sites on modified nucleosomes, places on the nucleosomes that are structurally more accessible than for the canonical and the modified nucleosomes. The arrows on the left (→) indicate DNase I 10 bp cuts on nucleosomes. The bar on the left represents the cERE (74-88 bp). 138

Figure 56. DNase I of tailless nucleosomes and N* (4E0). Lane 1, A/G ladder; lanes 2-4, nucleosomes digested for 30, 60 and 90 seconds, respectively; lanes 5-7, modified nucleosomes digested for 30, 60 and 90 seconds, respectively. The reaction was performed at 4 oC to maintain the integrity of the modified nucleosomes. The arrows on the right of the picture (←) indicate additional DNase I cut sites on modified nucleosomes, places on the nucleosomes that are structurally more accessible than for the canonical and the modified nucleosomes. The arrows on the left (→) indicate DNase I 10 bp cuts on nucleosomes. The bar on the left represents the cERE (114-128 bp).

139

Size of hypersensitive and new bands in (bp) DNA N’/N” N* 2E2 56, 74, 92, 97 and 109 50, 51, 52, 59, 60, 61, 62, 71, 82, 83, 92 and 98 3E1 70, 90, 92,120, 125, 132 and 140 4E0 50, 51, 52, 61, 62, 71, 79, 50, 51, 52, 59, 60, 61, 82, 93, 94 and 102 83, 84, 104, 112 and 140

Table 8. Location of additional DNA bands in N’/N” and N* from the DNase I experiment.

The table shows the appearance of 5 bands for 2E2 N’/N” and 12 bands in N*. There are 7 bands observed N’/N” for 3E1. There are 10 bands for 4E0 N’/N” and 12 bands in N*.

140

Figure 57. Comparison of additional DNase I sensitive cuts in the tailed (N’/N”) and tailless (N*) nucleosomes for the three DNA constructs. The scheme shows the size of additional DNase I bands in N’/N” (above the line) and N* (below the line). For the tailed nucleosomes, there are 5 bands (56, 74, 92, 97 and 109 bp) for 2E2 N’/N”, 7 bands (70, 90,

92,120, 125, 132 and 140 bp) for 3E1 N’/N” and 10 additional bands (50, 51, 52, 61, 62,

71, 79, 93, 94 and 102 bp) for 4E0 N’/N”. For the tailless nucleosomes, there are 12 bands observed in 2E2 N* (50, 51, 52, 59, 60, 61, 62, 71, 82, 83, 92 and 98 bp) and 12 observed bands in (50, 51, 52, 59, 60, 61, 82, 83, 84, 104, 112 and 140 bp) 4E0 N*. The dashed box represents 60 bp of DNA interaction with histone H3/H4 tetramer in the nucleosome.

Thirty base pair to either side of the dashed box interacts with H2A/H2B while the last 13 bp at the entry and exit point of the nucleosome is organized exclusive by the α-helical

histone fold extension of H3 and the preceding H3 N-terminal tail (Luger and Richmond,

1998). 141

27. Ava I digestion of tailless nucleosomes and modified tailless nucleosomes

The 161 bp DNA used in these experiments was designed to have an Ava I site at different positions on the DNA (Ava I site; 2E2 is 98 bp from the EcoRI end of the DNA,

4E0 is 138 bp from the end of the DNA). To further determine if HMGB1 changes the

character of tailless nucleosomes, the accessibility to Ava I sites were determined for

tailless and modified tailless nucleosomes. Identical number of counts of radiolabeled

nucleosomes was incubated with 2 U of Ava I and the reaction stopped at various times.

The DNA was purified and run on 4% polyacrylamide gel.

We have previously shown that compared to the canonical nucleosomes (N), Ava I cuts a larger percentage of the nucleosomal DNA in the HMGB1-modified nucleosomes.

The same general results were observed with the tailless nucleosome. Ava I digestion was probed over a period of 30 minutes to determine if the removal of these tails facilitated increased cleavage activity. Figure 58 showed that after 5 minute, Ava I cleaved about 52% of the nucleosomal DNA in the tailless nucleosomes, compared to 73% in the modified tailless nucleosomes and only 24 % (figure 38) for nucleosomes. This suggests that Ava I has gained greater access to the tailless nucleosome compared to the canonical nucleosome and that the Ava I gained even greater access after HMGB1 effect to produce N*. After 30 minutes of incubation, 75 % of tailless nucleosomes were cleaved compared to 83% of N*.

This suggests that the cutting by Ava I is further enhanced in the N*. Table 9 shows a comparison of canonical nucleosomes with tailless nucleosomes. The data are consistent with the results that ER binds to the tailless nucleosomes even without the aid of HMGB1.

The ER binding site and Ava I site is more accessible in the modified tailless nucleosomes compared to any of the forms of tailed nucleosomes. 142

We suggest that the HMGB1 may have two functions. First, it helps to dissociate

the histone tails from the DNA, thereby exposing the DNA. Secondly, it also binds to the

minor groove of the DNA and decreases the extent of the histone/DNA interactions. An

increase in Ava I digestion was observed in N’/N” (figure 38), which suggests that the

HMGB1 helps to dissociate the histone tails and expose the DNA to Ava I. If the tails are

cleaved off as in the case of tailless nucleosome, the effect of HMGB1 becomes more

pronounced and the Ava I may rapidly and more extensively cleave the DNA. That is

consistent with why maximum digestion was observed in the HMGB1 remodeled tailless

nucleosomes.

Figure 59 A shows that Ava I was inaccessible to 2E2. After 90 minutes of

incubation, not a significant amount of DNA was cleaved by the Ava I, even in the

remodeled tailless nucleosomes, which was expected to give maximum cleavage. The

crystal structure of nucleosome does show the structure of DNA at the dyad axis being very

different from the rest of the DNA in the nucleosome. The uniform curvature of 30o per helical turn throughout most of the DNA in a nucleosome structure is kinked at the dyad axis (Battistini et al, 2010 and Luger et al, 1997). This may explain the inaccessibility of

Ava I to 2E2. Figure 45 B was performed at the same time, with 2E2 and 4E0 nucleosomes, to show a positive result. Figure 59 B shows that the Ava I was accessible to 4E0 nucleosomes but not 2E2 nucleosome. Since the reactions were performed under the same conditions, the only reason why 2E2 was not accessible to Ava I was due to the structure of the DNA at the Ava I restriction site where the kink in the DNA is found. 143

Figure 58. Ava I digestion of 4E0 tailless nucleosomes and remodeled tailless nucleosomes. Equal counts (cpm) of tailless nucleosome and remodeled tailless nucleosome were incubated in a 200 µL total volume, and digested with 100 U of enzymes.

Aliquots of 40 µL were taken at 0, 5, 10, 15 and 30 minutes into EDTA, final concentration 144

to 25 mM. The purified labeled DNA from the digested nucleosomes was run on 8%

denaturing gel for 90 minutes. Lanes 1-5, DNA from the tailless nucleosome; lanes 6-10,

remodeled tailless nucleosomes. B. Kinetic plot of the percent Ava I digestion versus time

for tailless nucleosomes and HMGB1-modified tailless nucleosomes. The plot shows the

time profile for the percent of DNA that was cut by Ava I to produce DNA fragment of 138

bp tailless nucleosomes (▲) and modified tailless nucleosomes (●). The percent digestion was calculated from phosphoimager analysis amount of DNA cut after the experiment as a function of total DNA.

Percent Digestion of 4E0 nucleosomes Time Tailless (minutes) Nucleosomes N’/N” Nucleosomes N* 0 0 0 0 0

5 24 60 52 73 10 31 67 59 77

15 35 70 63 79

30 41 75 68 83

Table 9. Percent digestion of 4E0 nucleosomes with Ava I. The table shows the amount of

DNA digested by Ava I at various times. The percent digestion was calculated by dividing

the amount of nucleosome digested by total nucleosomes. An average of two separate

experiments was used in the calculation.

145

Figure 59. Ava I digestion of 2E2 nucleosomes (remodeled tailed and tailless nucleosomes). Equal counts (cpm) of nucleosome and remodeled nucleosome were incubated in a 200 µL total volume, and digested with 100 U of enzymes. Aliquots of 40 146

µL were taken at 0, 15, 30, 60 and 90 minutes into EDTA, final concentration to 25 mM.

The purified labeled DNA from the digested nucleosomes was run on 8% denaturing gel for 90 minutes. (A) Lanes 1-5, DNA from the 2E2 nucleosomes; lanes 6-10, remodeled nucleosomes; lanes 11-15, tailless nucleosomes; lanes 16-20, remodeled tailless nucleosomes. (B) Lanes 1-5, 2E2 nucleosomes; lanes 6-10, 4E0 nucleosomes.

28. Effect of temperature on HMGB1-remodeled tailless nucleosomes (N*).

The thermal stability of tailless and remodeled tailless nucleosomes (4E0) was

determined by incubation at different temperatures. Aliquots of tailless remodeled

nucleosomes were incubated at 4 oC, 37 oC and 55 oC, and compared with tailless

nucleosomes incubated at 37 oC.

Figure 60 showed tailless nucleosomes (4E0) were stable at 37 oC, even for a

prolonged incubation time of overnight. Lanes 5-8 show that the remodeled nucleosomes

(N*) were stable overnight at 4 oC. However, the N* were unstable at 37 oC and 55 oC and completely revert to canonical tailless nucleosomes. However, in contrast to the effect of temperature on N’/N”, the tailless revert back to the stable nucleosome, while the tailed nucleosome remain in the remodeled form, N’ (figure 39). This suggests that the tails of the core histones in N’ and N” are somehow involved in the formation of the N’/N” structure, and in stabilizing the N’ population. 147

Figure 60. Effect of temperature on tailless nucleosomes and remodeled tailless nucleosomes (4E0). Lanes 1-4: nucleosome in TE/sucrose buffer at 37 oC for 0, 60, 120 minutes and overnight (O/N), respectively. Lanes 5-8, remodeled tailless nucleosomes in

TE/sucrose buffer at 4 oC for 0, 60 and 120 minutes and overnight (O/N), respectively.

Lanes 9-12, remodeled tailless nucleosomes in TE/sucrose buffer at 37 oC for 0, 60,120 minutes and overnight (O/N), respectively. Lanes 13-16, remodeled tailless nucleosomes in

TE/sucrose buffer at 55 oC for 0, 60 and 120 minutes and overnight (O/N), respectively.

After the incubation, a 10 µL of each of nucleosome or remodeled nucleosome were loaded onto an EMSA gel and run for 2 hours at 200V at 4 oC. 148

29. Effect of increasing salt concentration on HMGB1- remodeled tailless nucleosomes

The stability of the modified tailless nucleosomes in increasing NaCl concentration was determined by incubating modified nucleosomes with 0-300 mM NaCl. Figure 61 shows, that unlike heat, addition of NaCl converts N* back to the canonical nucleosomes at

100 mM NaCl. This indicates that remodeled tailless nucleosomes are increasingly less stable as the NaCl concentration increases and reverted to a position that is close to that for the canonical nucleosomes at NaCl concentration of 50-100 mM. There is little difference between the tailless and tailed remodeled nucleosome since both revert back to canonical nucleosomes at about 100 mM NaCl. 149

Figure 61. Effect of increasing NaCl concentration on mobility of remodeled tailless nucleosomes (4E0). Lane 1, DNA; Lane 2, tailless nucleosome in TE/sucrose buffer; Lane

3, remodeled tailless nucleosome in TE/sucrose buffer; Lanes 4-8, remodeled tailless nucleosomes in TE/sucrose buffer with mM NaCl as indicated. D (DNA), N (nucleosome) and N’ and N” (HMGB1-remodeled nucleosome). After the incubation on ice for 15 minutes, the salt concentration in the reaction mixture were diluted to 100 mM for solutions in lanes 7 and 8 with those in lanes 4 and 5 (25 and 50 nM) initially kept at the same

NaCl concentration. After incubation, 20 µL of each were loaded onto an EMSA gel and run for 2 hours at 200V at 4 oC. 150

30. Effect of increasing levels of unlabeled DNA on EMSA mobility of modified tailless nucleosomes

We tested the stability of the modified tailless nucleosomes in the presence of excess unlabeled 161 bp DNA. It has been shown that incubation of nucleosomes with excess DNA disrupts the nucleosomes and allows for the formation of free DNA, dissociated from the nucleosomes (Workman and Kingston, 1992; Juan et al, 1993). Bands corresponding to free labeled DNA may be expected if excess unlabeled DNA competed for the core histones and facilitated DNA dissociation from the tailless nucleosomes.

However, figure 62 shows that there was no evidence of DNA dissociating from the histones. The remodeled tailless nucleosomes are, however, unstable in the excess unlabeled DNA and revert back to the canonical nucleosomes. The tailless nucleosome behaves similarly to the tailed nucleosome since both reverts back to canonical nucleosome when excess DNA is added to the reaction. Therefore, the tails do not have an effect on the stability of the HMGB1-remodeled nucleosome competing with excess DNA. 151

Figure 62. Effect of increasing the levels of unlabeled 161 bp DNA on HMGB1

modified tailless nucleosome. The modified tailless nucleosomes were incubated with

increasing amounts of unlabeled DNA on ice for 30 minutes and run for 2 hours on 4% polyacrylamide gel. Lane 1, DNA; lane 2, tailless nucleosomes; lane 3, remodeled tailless nucleosome from sucrose gradient; lane 4-8, remodeled tailless nucleosome treated with increasing levels of unlabeled DNA. D indicate free DNA, N (nucleosome) and N’ and N”

(HMGB1-remodeled nucleosome). 152

31. Translational position of DNA remains unchanged in HMGB1-remodeled nucleosomes.

We asked whether HMGB1- remodeled nucleosomes exhibited an altered translational position of DNA on the nucleosome, and therefore, allow ER to bind to cERE.

Exonuclease III cuts single strand DNA from the end in the 3’ to 5’ direction. The enzyme pauses when it encounters a protein or nucleosomal proteins. If HMGB1 changes the translation of DNA within the nucleosome, we would expect to find that Exo III digest the nucleosomal DNA into shorter fragments compared to the untreated nucleosomes.

Exonuclease III digestion was therefore, performed on N’/N” and on modified tailless nucleosomes (N*). Purified DNA was run on 8% denaturing gel for 3 hours.

Exo III reaction with 4E0 DNA produced cleavage and a series of shorter DNA fragments in both nucleosomes and HMGB1 remodeled nucleosomes (figure 63). This indicates that the Exo III gained equal access to DNA in all the nucleosomes (both canonical and HMGB1-remodeled. In Figure 36, the high HMGB1 concentration (1600 nM) appeared to be inhibiting the activity of the Exo III enzymes. However, in the

HMGB1-remodeled nucleosomes, there is little or no HMGB1 with the nucleosomes.

Therefore, HMGB1 had no effect on the translation of DNA. A change in the translation of

DNA in the nucleosomes would have resulted in the exonuclease gaining more access to the DNA in N’/N” or N*. Therefore, the HMGB1 does not alter the translational position of

DNA on the nucleosomes. 153

Figure 63. Exo III digestion on nucleosomes, HMGB1-remodeled nucleosomes, tailless

nucleosome and HMGB1-remodeled tailless nucleosomes. Nucleosomes were incubated

with or without 1600 nM HMGB1 on ice for 1 hour and remodeled nucleosome separated

on 5-30% TE/sucrose. The reaction was performed on ice to maintain the integrity of the

HMGB1-remodeled nucleosomes and digested with 1U/µL of Exo III. The reaction was

stopped at 1, 2 and 4 minutes. The DNA fragments were purified with phenol chloroform and ethanol precipitated. An 8% denaturing gel was run for 3 hours at 1500V. Lane 1, 5, 9 and 13, 161 bp DNA; lanes 2, 3 and 4; 6, 7 and 8; 10, 11 and 12 and 14, 15 and 16 are digestion stopped after1, 2 and 4 minutes, respectively.

154

32. DNase I digestion of tailless nucleosomes in the presence of 400 nM HMGB1

We have already shown that HMGB1 changes the character of nucleosomes by reducing its gel mobility and to enhance the binding of ER. We have also shown that a

DNase I pattern shows no difference between nucleosomes and nucleosomes treated with

400 nM HMGB1. To further characterize the effect of HMGB1 on tailless nucleosomes,

DNase I digestion was performed on tailless in the presence of 400 nM HMGB1. The suggestion is that the HMGB1 does two things; removes the tails from protecting the DNA, in addition to disrupting the interactions between the histones and DNA at higher HMGB1 concentration. With the tails removed, the HMGB1 can act to disrupt the histone/DNA interaction. We may anticipate that the DNase I pattern will show a difference between the tailless nucleosomes and the tailless nucleosomes treated with 400 nM HMGB1.

Figure 64 shows the presence of addition DNase I bands in the presence of 400 nM

HMGB1. This clearly indicates that in the absence of the tails, the 400 nM HMGB1 was sufficient to cause a disruption in the histone/DNA interaction.

155

Figure 64. DNase I of tailless nucleosome (4E0) in the absence and presence of 400 nM

HMGB1. Lanes 1-4, tailless nucleosomes digested for 30, 60, 90 and 120 seconds, respectively. Lanes 5-7, tailless nucleosomes treated 400 nM HMGB1 and digested for 30,

60, 90 and 120 seconds, respectively. The arrows on the right of the picture (←) indicate additional DNase I cut sites on modified nucleosomes, places on the nucleosomes that are structurally more accessible than for the canonical and the modified nucleosomes. The arrows on the left (→) indicate DNase I 10 bp cuts on nucleosomes. The bar on the left represents the cERE (114-128 bp).

156

CHAPTER 4: DISCUSSION

1. Characteristics of the rotationally phased and translationally positioned nucleosome

The nucleosome is the fundamental repeating unit of chromatin (Luger &

Richmond, 1998; Segal et al, 2006). A nucleosome packs about 147 bp of DNA wrapped

around a histone octamer (two copies of each of H2A, H2B H3 and H4) in 1.65 turns of a

left-handed superhelix and has a molecular weight of 210 kDa (Battistini et al, 2010;

Luger & Richmond, 1998). There is a curvature of ca. 30o per helical turn throughout most of the DNA in a nucleosome structure except for two sharper kinks of 50o at the

dyad axis (Battistini et al, 2010). All nucleosomes have a 10 bp per turn with the minor

groove of the DNA interacting with a histone at twelve independent locations. The 12

histone protein/DNA interactions are between the minor groove of the DNA and the

L1L2 loops & α1α2 DNA-binding motif on the histones (Luger, 2006, Luger et al, 1997).

DNA sequences differ greatly in the degree of difficulty to bend the DNA into

almost two complete turns around the histone octamer (Segal et al, 2006; Battistini et al,

2010). This process requires substantial energy to bend the DNA, primarily by

compressing the minor groove of the DNA helix. The repeating motif containing [5’-

(A/T)3 NN(G/C)3 NN-3’] has been shown to have a high affinity for binding the histone octamer and bending to accommodate a strong DNA-histone interactions in a nucleosome

(Shrader & Crothers, 1989). The sequence is termed the nucleosome positioning sequence (NPS). This sequence was used in the preparation of the 161 bp DNA in this project. There are four NPS in the 161 bp DNA construct flanking the cERE (M&M, box

1&2), which locks the cERE into a fixed rotational phasing and translational position. 157

The nucleosomes we used were prepared with 161 bp labeled DNA through the salt dilution method described in M&M, using oligonucleosomes prepared by partial MN digestion of chromatin from chicken erythrocytes. The oligonucleosomes were reacted with radiolabeled DNA at high salt concentration of 1 M and then diluted to 0.13 M. The nucleosome population was isolated by sedimentation on 5-30% sucrose gradient and run on 4% polyacrylamide gel. A single band representing the nucleosome was obtained

(figure 6). This is consistent with previous studies (Li et al, 1995; Workman et al, 1992 and Lorch et al, 1998) in which single band corresponding to the nucleosome was shown.

Since the nucleosome sediments at a different density compared to DNA, this procedure allows for a clean fractionation of the nucleosome from free uncomplexed DNA.

Exonuclease III digests DNA from 3’ to 5’end. The enzyme pauses on DNA if it encounters a bound protein or nucleosomal proteins. Exo III reaction was used to determine if nucleosome had the same translation and therefore, a homogenous population. Figure 46 indicates that the first induced stop was about 160 bp which is essentially the length of DNA we used. Such an exonuclease III digestion with the nucleosomes indicates that the nucleosomes population was homogenous.

The cERE was translationally and rotationally phased using NPS. In a rotationally phased nucleosome, the DNase I cuts in the outward facing, exposed minor groove every

10 bp (Imbalzano et al, 1994). DNase I digestion on nucleosomes show a 10 bp pattern, indicative of its rotational phasing (fig 13).

Binding studies with estrogen receptor(ER) indicates that the ER binds strongly to cERE in naked DNA with KD of 5 nM. However, incorporation of the cERE into nucleosomes drastically reduces the binding affinity of ER. We estimate that the KD of 158

ER binding to nucleosome, to be about 250-300 nM (fig 10). In all eukaryotes, the

genome is packaged in nucleosomes. The binding affinities of transcription factors are

reduced drastically when the binding sites are placed in a nucleosome (Chen et al, 1994,

Taylor et al, 1991, Sarpong, 2006). TBP binding to nucleosomes is drastically reduced by

105 (Imbazano et al, 1996). The binding of GAL4 and NF-1 to nucleosomes is reduced

by at least 100 fold (Chen et al, 1994, Taylor et al, 1991, Blomquist et al, 1999).

However, an exception is the glucocorticoid receptor (GR), which binds to a nucleosome

reconstituted on the promoter of the mouse mammary tumor virus with a 2-5 fold lower

affinity than to free DNA (Wrange, 1995). NF-kβ also binds to nucleosomes and DNA

with the same binding affinities (Angelov et al, 2004).

Before reactions with HMGB1, the normal nucleosomes are characterized as

stable in high salt buffer, heat or excess DNA and do not dissociate into free DNA and

histone octamers under these conditions (Workman and Kingston, 1992).

2. HMBG1 enhances ER binding to cERE within nucleosomes

HMGB1 is a very mobile “architectural” protein that binds nonspecifically in the minor groove of DNA (Bustin, 1999; Agresti & Bianchi, 2003) and has been shown to enhance transcription of a subset of genes by RNA pol II (Agresti & Bianchi, 2003; Das et al, 2004). HMGB1 and a related protein HMGB2 are reported to enhance binding of estrogen, glucocorticoid and androgen receptors to their recognition elements but have no effect on non-steroidal nuclear receptors (Boonyaratanakornkit et al, 1998). Our lab has shown that HMGB1 protein enhances the binding of estrogen receptor (ER) to a variety 159

of estrogen response elements (ERE) (Das et al, 2004; El Marzouk et al, 2008;

Ghattemani, 2004; Sarpong, 2006), with KD decreased from 5 nM to 2 nM.

In this work, we find that ER binds to 161 bp DNA with a KD of about 5 nM

(figure 7). However, the binding affinity is reduced if the same DNA is incorporated into

a nucleosome. The KD for ER binding to a nucleosome increases from 5 nM in DNA to

over 250-300 nM in the nucleosome (figure 20). Therefore, ER binding to cERE in

nucleosomal DNA is reduced ca 50-60 fold from that in DNA.

The presence of 400 nM HMGB1 facilitates ER binding within nucleosomal

DNA. The binding affinity of ER binding to 2E2 or 4E0 in the presence of 400 nM

HMGB1 was 50 nM. The binding is independent of the translational position of the

cERE in the 161 bp DNA construct. This is similar to NF-kB binding to nucleosomes, in

which NF-kB binding affinity is independent of the translational position of the binding

site on the nucleosomes (Angelov et al, 2004). Therefore, the presence of 400 nM

HMGB1 enhanced ER/cERE binding by at least 5-6 fold.

DNase I footprint (figure 24) shows that ER binds in a sequence-specific manner to cERE in a nucleosome. The mechanism by which HMGB1 enhances binding is not clear but there have been mechanisms suggested as to how this may be possible.

HMGB1 could make the cERE site in the nucleosome more accessible for ER binding by interaction of the acidic domain of HMGB1 with the positively charged histone tail domains removing them from the DNA surface and therefore, and exposing the DNA. A second possibility is that HMGB1 interacts with the ER itself to enhance binding to a nucleosome. Using GST-pull down assay, it was shown that the CTE of ER DBD interacts with HMGB1 to enhance ER binding to DNA (Melvin et al, 2004). It has also 160

been suggested that HMGB1 binds to and widens the minor groove to permit the C-

terminal extension (CTE) to directly interact with bps to enhance further binding (Scovell and Das, 2001). HMGB1 may also act as a link between the CTE and a binding site in the minor groove of the DNA (Romaine et al, 1998; Edwards, 1998).

The HMGB1 had the greater effect on ER binding affinity when the cERE was incorporated into a nucleosome, compared to naked DNA. The KD for ER binding in free

DNA was 5 nM and decreased to 2 nM in the presence of 400 nM HMGB1, which

represent a 2 fold increase in binding affinity. In the nucleosome, the binding affinity changed from 250-300 nM in the absence of HMGB1 to 50 nM in the presence of 400 nM HMGB1. Therefore, since the HMGB1 effect is significantly greater in ER binding to

nucleosomal DNA (factor of 5-7-fold), it is more plausible that the primary effect of

HMGB1 is to change the character of the nucleosomes to enhance ER binding.

The Nhp6 protein, which is a component of the yeast FACT protein, also

enhances the binding of transcription factor (Pob 3) to nucleosomes, albeit at high Nhp6

levels of 10 µM. HMGB1, on the other hand, allows for binding of estrogen receptor at a

much lower levels of 400 nM. This suggests that perhaps, the HMGB1 is much more

effective (at lower levels) in changing the nucleosome/chromatin structure to allow for

the binding of a transcription factor.

DNA ligase I carries out the final step in sealing gaps in DNA during replication

and DNA repair. DNA ligase I was found to access DNA that is wrapped about the

surface of a nucleosome in vitro and carries out its enzymatic function without help from

any of the chromatin remodeling factors (Chafin et al, 2000). It appears that ER and 161

many transcription factors require additional help to gain access and bind to their

recognition sites within the nucleosome.

3. Increasing levels of HMGB1 affect EMSA mobility of nucleosomes

To determine the effect of HMGB1 on nucleosome mobility and structure, nucleosomes were incubated with 400, 800 or 1600 nM HMGB1 for one hour at 4 oC and the EMSA mobility determined (figure 26). The mobility of the nucleosome bands steadily decreased as the concentration of HMGB1 was increased from 400 to 1600 nM.

The reduced mobility could not be due to stable binding of HMGB1 to the nucleosomes because previous work showed that HMGB1 did not form a stable component of the newly formed complex, as reflected in no supershift of the bands on addition of anti-

HMGB1 (Sarpong, 2006). A possible effect might be that HMGB1 acidic tails could be interacting with the histone tail domain and dissociating the tails away from the DNA, therefore exposing the DNA. We can also infer that HMGB1 does not block access to specific sites in the nucleosomal DNA or prevent the binding of transcription factors to the nucleosome.

The HMGB1 remodeled nucleosome has a lower mobility on EMSA gel compared to canonical nucleosomes. Since the charge of the nucleosome remains the same, the only variable that may be different is the size or shape of the nucleosome. This may be consistent with localized loss of interaction between the histones and the DNA, which would perhaps increase the overall size of the HMGB1-remodeled nucleosome. 162

4. HMGB1 remodels nucleosomes into a complex of lower EMSA mobility (N’/N” or HMGB1-remodeled nucleosomes)

To further characterize the nucleosomes in the lower mobility band formed in the presence of high levels of HMGB1, an attempt was made to isolate it using sedimentation in a linear sucrose gradient. Nucleosomes were treated with 1600 nM HMGB1 for an hour on ice and the reaction mixture sedimented on 5-30% TE/sucrose gradient. The fractions collected showed virtually the same mobility (slightly lower) as the normal nucleosome (i.e., no change in sedimentation coefficient), while EMSA revealed two new populations of HMGB1-remodeled nucleosomes (HR-NS) which were designated N’ &

N” and have distinctly different mobilities compared to that of canonical nucleosome (N)

(figure 29). Since HMGB1 is not stably bound to the nucleosome and has a much smaller sedimentation coefficient, the N’/N” complexes are in an environment with little or no

HMGB1. The molecular weight of HMGB1 is 25 kDa and that of a nucleosome is about

200 kDa. N’/N” are stable over two week period. Joshi (2010) has previously isolated

N’/N” and using sedimentation analysis and atomic force microscopy found no evidence of formation of a dinucleosome as previously suggested for a remodeled nucleosomes

(Schnitzler et al, 1998). Our DNA was constructed with four nucleosome positioning sequences which lock the DNA into a fixed rotation and translational position.

Supershift assays with antibodies to the individual core histones indicated that the

N’/N” contained all the 4 core histones (figure 30), and therefore, the N’/N” complex is not due to histone loss from the nucleosome complex. Reaction of N’/N” with anti-

HMGB1 had little or no effect on the mobility of N’/N” indicating that the remodeled 163

nucleosome, once formed, did not require HMGB1 to maintain its remodeled structure

(fig 44).

We also find that N’/N” remodeled nucleosomes were able to bind to ER without

the addition of any additional HMGB1, with a KD of 20 nM (fig 31). However, the

addition of poly (dI-dC) to the ER binding buffer causes the remodeled nucleosome to revert to canonical nucleosomes but still permit ER to bind to the nucleosomes with a

KD of ~ 40 nM, indicative of weaker binding of ER (fig 31). The poly (dI-dC) reduces some of the nonspecific binding of the ER to the modified nucleosome. However, the omission of the poly (dI-dC) in ER dilution buffer had no effect on ER binding to canonical nucleosomes (data not shown).

The instability of the remodeled nucleosome in the ER dilution buffer (final buffer concentration; 40 mM KCl, 5%(w/v) glycerol, 7.5 mM Tris-HCl, pH 8.0, 0.1 mM

EDTA, 0.2 mM DTT, 50 ng/µL BSA, 1 ng/µL poly (dI-dC) led us to examine whether the high salt component of the buffer was partially responsible for the observed effect.

N’/N” was incubated with increasing NaCl (figure 41).The remodeled nucleosomes were unstable as the salt concentration increased and reverted to canonical nucleosome at 50 mM NaCl. Further addition of NaCl (up to 300 mM) did not have any effect on the canonical nucleosome. N’/N” appears to be thermally and kinetically unstable. The

canonical nucleosome form is therefore, the most thermodynamically stable form.

A similar study on the effect of salt on altered nucleosome structures with RSC complex indicates that RSC complex can convert nucleosome in the presence of ATP to a slower migrating form (Lorch et al, 1998). The slow migrating form was however, unstable in high ionic salt environment, and converts into canonical nucleosomes (Lorch 164

et al, 1998). Although the altered nucleosomes are stable in low salt buffer without

HMGB1, it appears plausible that HMGB1 is required in high salt buffer to produce and

maintain the nucleosome in a remodeled state.

The thermal stability of the HMGB1-remodeled nucleosomes was also examined

by incubation at increasing temperatures (fig 40). We find the canonical nucleosome

form to be the most thermally stable form. N” reverts to N’ with no evidence of canonical nucleosome being formed even on overnight incubation at 37 oC and 55 oC.

Competition experiment with excess unlabeled DNA showed that the HMGB1-

remodeled nucleosomes were unstable in excess DNA, and reverted to the canonical

nucleosomes. However, there was no evidence of the formation of free labeled DNA as

previously observed (Workman & Kingston, 1992). Kingston’s group presented data to

show that incubation of nucleosomes with excess unlabeled DNA disrupted nucleosomes and allowed free labeled DNA to dissociate from the nucleosomes. The complex formed when GAL4 binds to a nucleosome (containing five GAL4 binding sites) was unstable in excess nonspecific competitor DNA (without GAL 4 binding site), and revert to either the original nucleosome core particle or GAL4 bound to naked labeled DNA (Workman &

Kingston, 1992). Bands corresponding to free labeled DNA may be expected if excess unlabeled DNA competed for the core histones and facilitated DNA dissociation from the nucleosomes. However, fig 41 showed that there was no evidence of DNA dissociating from the HMGB1-remodeled nucleosomes.

Overall, HMGB1 is able to remodel the nucleosomes into a different structure through a mechanism that does not require ATP or ATP hydrolysis. The HMGB1- remodeled nucleosomes are trapped in a higher energy state that is unstable in high NaCl, 165

excess DNA and heat. The HMGB1-remodeled nucleosome does not require HMGB1 to

maintain its conformation (fig 44) and, is stable for at least a month at – 20 oC, compared to nucleosomes remodeled by SWI/SNF which revert back to canonical nucleosomes within 30 minutes after ATP removal (Imbalzano et al, 1996).

5. HMGB1 can alter the DNase I 10 bp digestion pattern within a remodeled

nucleosome

DNase I can be used to define the rotational phasing of the DNA within a

nucleosome (Imbalzano et al, 1994). For rotationally phased nucleosome, the DNase I

cuts in the exposed minor groove every 10 bp (with some being more intense than

others). If the phasing of the DNA within the nucleosomes is altered, the 10 bp pattern

will be changed, usually with the addition of more bands, and a pattern approaching that

for free DNA (Imbalzano et al, 1994).

DNase I digestion on nucleosomes in the presence of 400 nM HMGB1that

permits ER binding, had a pattern identical to that for the canonical nucleosomes (figure

25). Therefore, although 400 nM HMGB1 facilitated ER binding, it was not sufficient to

alter the rotational position of DNA in the nucleosomes to an extent that could be detected on the denaturing gel. On the other hand, DNase I digestion of nucleosome in

the presence of 1600 nM HMGB1 (Joshi, 2009) or on HMGB1 remodeled nucleosomes

(N’/N”) did show the presence of additional DNase I cuts while maintaining the basic 10

bp pattern. Figures 33, 34 & 35 show that the remodeled nucleosomes exhibit additional

bands compared to the canonical nucleosomes. Those additional sites on the remodeled

nucleosome reveal an increased sensitivity to DNase I cutting which was not observed in 166

the canonical nucleosomes. This suggests that the structures of HMGB1-remodeled

nucleosomes (N’/N”) are different than the nucleosome and minor grooves have an

increased and broader accessibility at those points that show DNase I cutting sensitivity.

The addition DNase I cuts span the entire length of the 161 bp DNA construct used to

prepare the nucleosomes (56-140 bp).

The Saccharomyces cerevisiae yFACT complex consist of three proteins; Nhp6,

Pob3 and Spt16 (Formosa et al, 2001). The Nhp6 unit contains a single DNA binding

motif similar to those found in members of the HMG box family of DNA binding proteins (Ruone et al, 2003). yFACT has been shown to mildly enhance the sensitivity of specific sites within nucleosomal DNA to DNase I and also, alter nucleosome structure in a manner that does not require ATP hydrolysis (Formosa et al, 2001). Nhp6 protein, which is only one component of yFACT, can independently convert a canonical nucleosome into another form that is more accessible to DNase I cleavage at specific sites

(Rhoades et al, 2004). Our results are similar to those of Nhp6 because HMGB1- remodeled nucleosomes were more accessible to DNase I compared to canonical nucleosomes. A point where we differ is the amount of protein needed to remodel the canonical nucleosomes. To remodeled nucleosomes, Formosa’s group used 10 µM of

Nhp6 which is 10-fold higher in Nhp6 concentration than the 1600 nM HMGB1 we used

(Ruone et al, 2003). The suggestion is that the mechanism of action for HMGB1 may include; 1) interacts with the histone tail domain and lead to greater dissociation from the nucleosome; 2) global change in the nucleosome structure by loosening the histone/DNA interactions or perhaps an increase in the flexure of DNA (Travers, 2003). Essentially, at

400 nM of HMGB1, we think that HMGB1 binds the histone tail domains without 167

disrupting the interaction between the DNA and the core histone loops. This is consistent

with the unchanged 10 bp pattern observed in the presence of 400 nM HMGB1 where no

additional DNase I sensitive bands were observed (fig 13). At 1600 nM, however, the

higher level of HMGB1 has the additional effect of globally disrupting the interaction

between the globular regions of the core histone and the DNA. The nucleosome becomes

more dynamic and changes, as revealed by DNase I activity in regions beyond the 10 bp

site, therefore accounting for the additional bands observed in the HMGB1-remodeled

nucleosomes.

Other known chromatin remodeling multisubunit complexes such as SWI/SNF

and imitation switch (ISWI) are ATP-dependent. These complexes can work by both

altering DNA/histone interaction in addition to translocation of DNA relative to the

histone proteins (Imbalzano et al, 1996; Phelan et al, 2000). SWI/SNF alters the rotational phasing of DNA in nucleosome to facilitate TBP binding to the TATA box in a nucleosome (Imbalzano et al, 1994). RSC, a SWI/SNF related chromatin remodeling complex, is also ATP-dependent (Lorch et al, 1998). HMGB1 on the other hand, is not an ATPase and does not translocate DNA. The remodeling process by HMGB1 is ATP- independent. RNA polymerase can not initiate transcription on nucleosomal DNA in vitro

(Lorch et al, 1987), and there is the need for a localized disruption of chromatin (Pham et al, 1991) before transcription can start. We suggest that, these proteins, acting in concert with HMGB1, may be involved in disrupting the nucleosome before regulatory factors such as ER can bind and before RNA polymerase can initiate transcription. 168

6. Exonuclease III shows that HMGB1 does not affect the translation of DNA in

HMGB1 remodeled nucleosomes

Exonuclease III acts on double stranded DNA by cutting single strand DNA from

an end in the 3’ to 5’ direction. The enzyme pauses when it encounters a protein or nucleosomal proteins. If the action of HMGB1 changes the translation of DNA within the nucleosome, we would expect to find that Exo III digests the nucleosomal DNA into shorter pieces compared to the untreated nucleosomes.

HMGB1 does not increase the access of Exo III to the ends of the DNA in the

N’/N” (fig 63). The Exo III experiment with 1600 nM HMGB1 was inhibitory to the Exo

III enzyme (fig 36). However, the N’/N” was effectively devoid of HMGB1, and showed the same Exo III digestion profile as observed on nucleosomes (i.e., no change). This suggests that the DNA near the exit/entry points of the nucleosome remain similarly bound to the nucleosome and did not unwind and dissociate as suggested by site exposure theory (Anderson et al, 2002). HMGB1 therefore, does not alter the position of the

histone octamer in relation to the DNA.

yFACT also remodels nucleosomes and likewise does not alter the position of

nucleosome indicating that it also acts through a mechanism distinct from ATP-

dependent DNA translocation and perhaps similar to that of HMGB1(Rhoades et al,

2004). yFACT only enhances access to Exo III at a very high enzyme levels of 0.2 µM

(Rhoades et al, 2004).

HMGB1 has been shown to bind randomly to the minor groove DNA to cause a

local distortion in the double helix (Bustin, 1999; Wang et al, 2007, Muller et al, 2001).

The minor groove of DNA interacts at twelve independent locations on a nucleosome, 169

with the helix-loops on the histones (Luger et al, 1997; Luger and Richmond, 1998a;

Luger, 2003). We think that, there may be a competition between the L1& L2 loops and

α1 & α2 DNA-binding motif of histone tails and HMGB1 for the minor groove in the

DNA (figs 3 & 63). Perhaps the HMGB1 interferes with the interaction between the L1&

L2 loops and α1 & α2 DNA-binding motif of the histone tails and the DNA, therefore,

allowing a local detachment of the DNA from the histones. This may allow for the

formation of a transient detachment between the DNA and the histone core of the

nucleosomes (as proposed by Travers, 2003). Assuming that is true, the transient

detachment formed may increase access of DNase I to sites in the N’/N” and change the

DNase I 10 bp pattern and also allow the binding of ER to the nucleosome.

7. HMGB1 enhances accessibility of Ava I to HMGB1-remodeled nucleosomes

Experiments on the effect of HMGB1 on access in the nucleosome was also

investigated via the effect on Ava I cutting within nucleosome. Ava I was located at 98

(2E2) and 138 (4E0) from the labeled end of the DNA. The suggestion of transient

detachment is also consistent with the data on Ava I digestion of nucleosomes. The

restriction enzyme Ava I has one recognition sequence within each of the DNA construct

(fig 4). A measure of accessibility of the site was determined by measuring the extent to

which Ava I cleave the DNA within the nucleosome. The data show that N’/N” were

more accessible to Ava I digestion compared to canonical nucleosomes (fig 39). The

initial rate of digestion at (0-5 minutes) for 4E0 nucleosomes was 5%/min and 12%/min

for N’/N”. This suggest that the structure of N’/N” is very different and more accessible 170

compared to canonical nucleosomes. All the restriction enzyme experiments were performed with 0.5 U/ µL Ava I.

Nhp6 protein enhances some restriction enzyme digestion (Hha I but not Sty I)

(Rhoades et al, 2004). Our results compare with those of Nhp6 because HMGB1-

remodeled nucleosomes were more accessible to restriction enzyme digest.

There were two DNA constructs made with the restriction sites of Ava I placed at

two different translational positions (dyad (2E2) and 40 bp from the dyad (4E0). The 4E0

was the more accessible of the two constructs. After 30 minutes, 41% of total

nucleosomal DNA was cleaved compared to 75 % in the remodeled nucleosome.

Interestingly, the digestion of 2E2 was different. The 2E2 N’/N” was inaccessible to Ava

I even after 90 minnutes of incubation (figure 57). Less than 10% of total nucleosomes

were cleaved in all cases. This is not an isolated incident because it has been shown that

not all restriction endonuclease can gain equal access to sites on the same nucleosome, or

a change in the placement of the restriction site may change the accessibility of the

restriction enzyme (Anderson et al, 2002; Chafin et al, 2000 and Xin et al, 2009). The

crystal structure of nucleosome does show that the structure of DNA at the dyad axis is

very different from the rest of the DNA in the nucleosomes (Battistini et al, 2010 &

Luger et al, 1997). The curvature of 30o per helical turn throughout most of the DNA in a nucleosome structure is kinked at the dyad axis (Battistini et al, 2010 & Luger et al,

1997). This may explain the inaccessibility of Ava I to 2E2. It has been suggested that the DNA in the nucleosome dissociate slightly from the ends through transient site exposure (breathing), which can then be recognized and cut by the restriction enzyme.

Our results for 4E0 N’/N” is similar to the proposal of nucleosome “breathing” as the cut 171 site of Ava I is only 23 bp from the end of the DNA (Li and Widom, 2004; Anderson and

Widom, 2000).

The activities of HMGB1, Nhp6 and yFACT are summarized in tables 10, 11 &

12 respectively.

8. Characteristics of the rotationally phased and translational positioned tailless nucleosome

We know that DNA within the context of a nucleosome reduces the binding affinity of most transcription factors, and in some cases, block transcription initiation and elongation (Biswas et al, 2004; Ge & Roeder,1994; Imbalzano, 1998). What is controversial is whether the histone tails play any part in reducing the binding affinity or, in our case, are they involved in HMGB-dependent remodeling of nucleosomes? The histone tail domains are highly positively charged and located at the N-terminal of all the core histones, in addition to the C-terminal of histone H2A. The histone N-terminal tails are important for chromatin assembly, remodeling (through post transcriptional modification eg. acetylation) and stability of the nucleosome (Ujvari et al, 2008; Yang et al, 2006).

Tailless nucleosomes have been used by many labs as a model system for highly acetylated nucleosomes (Polach et al, 2000). Acetylation of each of the basic amino acids means the loss of one positive charge. Hyperacetylation therefore reduces the overall positive charge in the histone tail domains. Hayes’s group has shown that removal of the tail domains do not affect the stability of translational or rotational position of the 172 nucleosome (Dong et al, 1989, Hayes et al, 1991). Trypsinized nucleosome remained intact (Polach et al, 2000).

The DNase I 10 bp digestion pattern on the tailless nucleosomes show that the 10 bp pattern remains unaltered, indicative that the rotational phasing in the nucleosome is independent of the presence of the tail domains (figure 55). The tailless nucleosomes were constructed using the same 161 bp DNA as in nucleosome (with tails). We therefore, did not expect much to change in terms of the DNA rotational phasing.

Exonuclease III reactions were used to test whether the translational position of the DNA in tailless nucleosome (without HMGB1) was the same as nucleosome. Figure

61 shows that the first induced stop in tailless was about 160 bp and same as that in nucleosome with tails. This suggest that even though the tails were cut off using trypsin, the nucleosome remained intact with the same translational position (and rotational phasing) as nucleosomes with tails.

In the absence of HMGB1, binding studies with ER to tailless nucleosomes indicate strong ER binding to cERE (figure 48) with a KD of 50 nM. This is in sharp contrast to nucleosome with tails, in which the KD was estimated at greater than 250 nM.

This result is similar to that found for TFIIIA binding to tailless nucleosome. It was reported that binding of TFIIIA to 5S RNA gene in a nucleosome was greatly reduced.

However, cleavage of histone tails with trypsin enhanced binding of the TFIIA (Vitolo et al, 2004). The binding of ER to tailless nucleosomes is also similar to the action of DNA ligase in binding to canonical nucleosome with tails (Chafin et al, 2000). 173

In the absence of HMGB1, the tailless nucleosomes were found to be stable in

high salt buffer, heat or excess DNA and do not dissociate into free DNA, effectively the

same stability profile observed for canonical nucleosome with tails.

9. HMBG1 enhances ER binding to cERE within tailless nucleosomes

The removal of the histone tail domains by trypsin digestion enhances ER binding to nucleosomes even in the absence of HMGB1. The KD for the binding of ER to tailless

nucleosome was 50 nM compared to the lack of binding to the canonical nucleosome

(with tails) in the absence of HMGB1. The binding affinity of ER to tailless nucleosomes

is further enhanced by the presence of 400 nM HMGB1 to 25 nM (figure 48). Our results

are similar to that found by the Hayes group, which show that tailless nucleosomes or

hyperacetylation of the nucleosome enhances binding of TFIIIA binding to the 5S gene in

the nucleosomes (Yang et al, 2005 and Vitolo et al, 2004). Taken together, these results

show that cleavage of the tail domains from the nucleosome enhances binding of

transcription factor to nucleosome without the need for a remodeling protein.

10. HMGB1 remodels tailless nucleosomes into a complex of lower EMSA mobility

To further characterize the tailless nucleosomes, which exhibit a lower mobility

band in the presence of high levels of HMGB1, an attempt was made to isolate the

nucleosome from using a linear sucrose gradient. Tailless nucleosomes were treated with

1600 nM HMGB1 for an hour on ice and the reaction mixture sedimented on 5-30%

TE/sucrose gradient. 174

The fractions collected shows that HMGB1-remodeled tailless nucleosomes (N*) had a different EMSA mobility compared to canonical tailless nucleosome, even though both complexes exhibited virtually the same sedimentation characteristics (figure 52).

The N* is essentially devoid of HMGB1, as reflected in the treatment with α-HMGB1

(figure 44) but the complex was stable for a period of at least 30 days. This suggests that

HMGB1 is not required continuously to maintain the remodeled state of the nucleosome.

The stability of the remodeled tailless nucleosome was determined by incubating the tailless nucleosome in increasing concentration of NaCl. It was observed that N*, similar to what was observed in N’/N”, were also unstable and reverted to canonical nucleosome at 50 mM NaCl (figure 61). Further addition of salt (up to 300 mM NaCl) did not dissociate the N* and it remained as a canonical nucleosome. The tailless nucleosome form is therefore, the most stable form at physiological salt conditions. This suggests that the nucleosome can equilibrate between different conformational states until stabilized in a single conformation in low salt buffers.

The thermal stability of the remodeled tailless nucleosomes was determined by incubating HR-tNS at different temperatures (figure 60). Unlike the nucleosomes with tails, the remodeled tailless nucleosome reverted to canonical tailless nucleosomes on overnight incubation at 37 o C and 55 o C, while N’/N” did not. The canonical nucleosome form (with tails) is therefore, the most thermally stable form of the two complexes. This suggests that the tails are important in maintenance of thermal stability of the nucleosomes.

Competition experiment with excess unlabeled DNA showed that the remodeled tailless nucleosomes were unstable in the excess DNA, and reverted to the canonical 175

tailless nucleosomes. The results were comparable to the nucleosomes (with tails), which

suggests that the tails were not important in competing with excess DNA to maintain the

remodeled state of the nucleosomes.

11. HMGB1 alters the DNase I 10 digestion pattern within a remodeled tailless nucleosome

DNase I can be used to define the rotational phasing of the DNA within a nucleosome (Imbalzano et al, 1994). For rotationally phased nucleosome, the DNase I cuts in the exposed minor groove every 10 bp. If the phasing of the DNA within the nucleosomes is altered, the 10 bp pattern will be changed, usually with the addition of more bands, and the pattern approaches that for free DNA (Imbalzano et al, 1994). If the

DNA/histone interactions are altered, the DNase I digestion pattern will reveal additional bands.

Figure 55 shows that the 10 bp pattern is maintained after cleavage of the tail domains. This indicates cleavage of the tail domains does not affect the rotational setting of the nucleosome. On the other hand, the N* show the presence of additional DNase I sensitive cut sites. This indicates that the remodeled nucleosome structure is very different from the canonical nucleosome and HMGB1 is able to remodeled tailless nucleosome, which is similar to SWI/SNF remodeling of tailless nucleosome (Guyon et

a., 1999). SWI/SNF remodeling complex remodels nucleosome in an ATP-dependent manner and the DNase I pattern after the remodeling process tends to resemble that on

free DNA (Guyon et al, 1999). In another study where the tail domains of H2A/H2B,

H3/H4, or all tail domains were trypsinized, human RNA pol II was able to produce a

greater transversal of nucleosome (Ujvari et al, 2008). Transcription by yeast pol II 176 proceeded much farther into nucleosome in the absence of tail domains. This is suggestive a loosely packed chromatin structure, which allows for the progression of pol

II.

12. Exonuclease III shows that HMGB1 does not alter translational position of DNA in HMGB1 remodeled tailless nucleosomes

Nucleosome remodeling by ATP-dependent remodeling complexes such as nucleosome remodeling factor (NURF) alters chromatin structure by catalyzing nucleosome sliding, thereby exposing DNA sequence in the nucleosome (Hamiche et al,

2001). Removal of N-terminal tail of histone H2B has been shown to promote uncatalyzed nucleosome sliding (Hamiche et al, 2001). The sliding of the DNA from the core particle can be detected by monitoring Exo III digestion profile.

Exo III digestion of N* presented no evidence for DNA translocation (figure 63).

The first induced stop of the Exo III was about 160 bp which is essentially the length of the DNA.

In a related study, it was shown that removal of tails from nucleosomes assembled on DNA fragment containing Xenopus borealis somatic-type 5S RNA gene result in repositioning of nucleosome 10 bp along the DNA (Yang et al, 2006). No ATP- dependent CRC were used in this study. The binding of the TF TFIIIA binding to the 5S gene displaces 20 bp of DNA-histone interaction at the periphery of the nucleosomes

(Vitolo et al, 2004). However, we see no evidence of movement in the DNA relative to the histone octamer as a result of the activity of HMGB1. This is consistent with the lack of ATPase activity in HMGB1, while being present in NURF.

177

13. HMGB1 enhances accessibility and cutting activity of Ava I to HMGB1- remodeled nucleosomes

Ava I digestion indicated that the tailless nucleosomes were more accessible compared to the canonical nucleosomes for the 4E0 nucleosome DNA. The initial rate after 5 mins of digestion was 5%/min for nucleosomes and 10%/min for tailless nucleosomes. Likewise, the N* was the most accessible among all the remodeled nucleosome structures tested. After 30 minutes of incubation of Ava I with N*, 83% of

N* was cleaved compared to 75 % of N’/N”. The initial rate of digestion after 5 mins was

15%/min for N* compared to 12%/min in N’/N”. This indicates that the tails of the core histone play a role in interacting with the HMGB1.

We suggested earlier that the mechanism of action for HMGB1 was to interact first with the histone tail domain and then globally loosen the histone/DNA interaction and/or perhaps increase the flexure of DNA. Essentially, in the tailless nucleosomes, the

HMGB1 disrupts the interaction between the DNA and L1& L2 loops and α1 & α2 DNA- binding motif interactions of the core histone.

HMGB1 can most readily alter and disrupt DNA interaction with the positively charged tails. At higher concentration, HMGB1 can also disrupt DNA/core histone interaction. In the absence of tails on histones, HMGB1 targets the DNA/core histone interaction and can affect these interactions at lower concentration that found in the tailed nucleosomes.

We find, however, that removal of the core histone tail domains does not result in increase Ava I digestion where the Ava I site is near the dyad axis (2E2). This is consistent with the finding that cleavage of tail domains does not result in increase in 178

exposure of DNA near the nucleosome dyad (Chafin et al, 2000). One might have

expected to see an increase in Ava I digestion of tailless 2E2 HMGB1 remodeled

nucleosomes. However, that was not the case. This may be attributed to the kink of the

DNA at the dyad axis. Depending on buffer conditions and the type of restriction enzyme used, it has been shown that removal of the histone tails increases access of restriction enzyme to the nucleosome by 1.5 to 14 fold (Polach et al, 2000).

In summary we suggest that HMGB1, by a non-enzymatic mechanism, interacts

with the nucleosome to generate a new population of “nucleosomes of altered

conformation”. Thus, HMGB1 provides an alternate, ATP-independent mechanism by

which a subset of transcription factors can gain access to their recognition sites within a

nucleosome and/or HMGB1 may cooperate with chromatin remodeling complexes to

enhance their activity.

The crystal structure of the nucleosome indicates that the histone tail domain

emerges from the core of the nucleosome to the outside of the DNA, and may protect the

DNA from restriction enzyme digest or binding of transcription factors. We suggest that, essentially at 400 nM of HMGB1, the histone tail domain interact with the HMGB1, and the tails are moved away from the nucleosome. This action may expose the DNA to restriction enzyme digest and to binding of transcription factors, and is consistent with our data. As the concentration of HMGB1 is increased to 1600 nM, the effect of HMGB1 becomes more pronounced and leads to the formation of a completely different nucleosome (N’/N”). In the N’/N” nucleosomes, we suggests that the interaction between the L1& L2 loops and α1 & α2 DNA-binding motif and the minor groove of nucleosomal

DNA is loosened by the action of the HMGB1 interacting with the histone tail domains 179 and the minor groove in the DNA (as illustrated in figure 65). Therefore, the nucleosomes become even more accessible to transcription factors binding, DNase I and restriction enzyme digestions.

Cleavage of the histone tail domains with trypsin further exposes the DNA in a nucleosome. The tailless nucleosome essentially behaves like the canonical nucleosome in the presence of 400 nM HMGB1 in which the tail domain-DNA interactions have been disrupted and the tails are now more freely extended from the nucleosome surface. The tailless nucleosomes binds to ER in the absence of HMGB1 with a Kd of about 50 nM which is consistent with the Kd of ER binding to canonical nucleosomes in the presence of 400 nM HMGB1. The tailless nucleosomes were also accessible to AvaI digestion in the absence of HMGB1. The effect of HMGB1 in remodeling tailless nucleosome is further enhanced in the absence of the histone tail domains (N*). The N* was therefore, the most accessible to AvaI digestion and the rate of AvaI digestion was also the highest.

In comparison, we suggest that the canonical nucleosome has a more compact structure compared to the HMGB1-remodeled nucleosomes (with or without histone tail domains). The interaction between the minor grooves of DNA and histones are weak in the remodeled nucleosomes compared to canonical nucleosomes. Therefore, it is easier for transcription factors to bind to the remodeled nucleosomes. It is also more accessible to DNase I and AvaI enzyme digestion. 180

Figure 65. Interaction between the minor groove of DNA, L1L2 loops (β) and α1α2

DNA-binding motif (N). The interaction between the L1& L2 loops and α1 & α2 DNA- binding motif and the minor groove of nucleosomal DNA is loosened in HMGB1- remodeled nucleosomes. (Picture was adapted from Ramakrishna, 1997). 181

Comparison of activities for HMGB1, Nhp6, and yFACT

Table 10: Activities for HMGB1 Activities References HMGB1 Interacts with minor groove of DNA (Bustin, 1999) Bends and modifies DNA structure (Wolffe, 1994) Facilitates transcription of a subset of genes and (Waga et al., 1988, 1990), elongation by RNA pol II Agresti & Bianchi, 2003 Reversibly inhibits transcription initiation by forming ( Ge and Roeder, 1994;Das HMGB1-TBP-TATA and Scovell, 2001; Dasgupta and Scovell, 2003) Stimulates transcription by relaxation of chromatin (Ogawa et al., 1995) Reorganizes nucleosome structure and increases This dissertation; Joshi, 2010 DNase I sensitivity DNase I sensitivity was enhanced on the entire length of the DNA This dissertation; Joshi, 2010 Does not require ATP for nucleosome remodeling This dissertation Facilitates chromatin remodeling by ATP-depended (Bonaldi et al., 2002; chromatin remodeling complexes Ugrinova et al., 2009; Yamada et al., 2004) Facilitates maintenance of genome stability (Giavara et al., 2005)

Converts nucleosomes between canonical and less This dissertation stable forms Facilitates the assembly of RAG1/2 recombination signal sequences DNA complexes that are Lange and Vasquez, 2009; components for V(D)J cleavage. Swanson, 2004

During transcriptional activation of pS2 gene, H1 is Ju et al, 2006 replaced by HMGB1/2 in a nucleosome that has one ER binding site

182

Table 11: Activities for Nhp6 Activities References Nhp6 Interacts with minor groove of DNA (Allain et al., 1999) Bends and modifies DNA structure (Allain et al., 1999) Facilitates transcription initiation by pol III (Kassavetis and Steiner, 2006; Kruppa et al., 2001; Lopez et al., 2001) Facilitates assembly of TBP-TFIIA-TATA on (Biswas et al., 2004; DNA Eriksson et al., 2004) Facilitates in transcriptional initiation by pol II (Biswas et al., 2004; Eriksson et al., 2004) Reorganizes nucleosome structure and (Rhoades et al., 2004; increases DNase I sensitivity Ruone et al., 2003) Facilitates global accessibility of nucleosomal (Xin et al., 2009) DNA by hydroxyl radicals Does not require ATP for nucleosome (Xin et al., 2009) remodeling Facilitates maintenance of genome stability (Giavara et al., 2005) 183

Table 12: Activities for yFACT Activities References yFACT Contains Nhp6 and interacts with DNA in the (Formosa et al., 2001) nucleosome Facilitates transcription elongation by pol II (Formosa et al., 2001; Formosa et al., 2002; Krogan et al., 2002; Mason and Struhl, 2003; Orphanides et al., 1998; Saunders et al., 2003) Stimulates transcription initiation by facilitating (Biswas et al., 2005; Mason TBP and TFIIA binding to nucleosomal sites and Struhl, 2003) Interacts with DNA polymerase α, and with (Gambus et al., 2006; MCM proteins, and it plays an important role in VanDemark et al., 2006; DNA replication Wittmeyer et al., 1999) Reorganizes nucleosome structure and increases (Rhoades et al., 2004; DNase I sensitivity Ruone et al., 2003) DNase sensitivity was enhances at the sites (Rhoades et al., 2004; clustered near the center of the nucleosomal DNA Ruone et al., 2003) away from entry and exit points. Facilitates global accessibility of nucleosomal (Xin et al., 2009) DNA by hydroxyl radicals Does not require ATP for nucleosome remodeling (Rhoades et al., 2004; Ruone et al., 2003; Xin et al., 2009) Converts nucleosomes between canonical and less (Xin et al., 2009) stable forms

184

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Appendix

1. Presence of ATP has no effect on binding of ER to nucleosome in the presence of 400 nM HMGB1.

The binding of ER to nucleosomes was performed in the absence and presence of

4 mM ATP to determine if ATP had any effect on ER binding in the presence of 400 nM

HMGB1. Appendix 1 shows that HMGB1 enhances the binding of ER to nucleosomes.

However, we do not observe an influence on the binding of ER to nucleosomes in the presence of 4 mM ATP. In the presence of both ATP and HMGB1, the binding affinity

ER to nucleosomes was not enhanced. This suggests that the binding of ER to nucleosomes in the presence of 400 nM HMGB1 is ATP-independent. 195

Appendix 1. Effect of ATP on ER binding to nucleosomes in the presence of 400 nM

HMGB1. Nucleosomes were incubated on ice with increasing concentration of ER, in the presence or absence of 4 mM ATP, for 30 minutes. Lanes 1-4 shows ER binding to a

nucleosome in the presence of 4 mM ATP, lanes 5-8 in the presence of 400 nM HMGB1

and lanes 9-12 in the presence of both 4 mM ATP and 400 nM HMGB1. ER

concentrations are 0, 25, 50 and 100 nM. 196

2. Binding of ER to nucleosomes in the presence of 400 nM HMGB1

We examined the ER binding to cERE in a nucleosomal DNA in the presence of

400 nM HMGB1. Appendix 2a, 3a and 4a show that ER binds to nucleosomes in the presence of 400 nM HMGB1. The binding has been shown previously to be independent of the position (2E2, 3E1 or 4E0) of the ERE on the nucleosomes, with a KD of about 50 nM (Sarpong, 2006). Appendix 2b, 3b, 4b shows the binding profile of ER binding to nucleosomes in the presence of 400 nM HMGB1, with a KD of 50 nM. The graph was plotted dividing the amount of complex formed by the total amount of DNA (M&M). 197

B 100

% Complex 20

1 10 100 ER [nM] 198

Appendix 2. ER binding to 2E2 nucleosomes in the presence of 400 nM HMGB1.

Nucleosomes were incubated for 30 mins with increasing concentration of ER on ice, and run on 4% polyacrylamide gel for 2 hours. Lanes 1-8 represents ER concentrations 0, 10,

20, 30, 40, 50, 60 & 80 nM, respectively. ER binds significantly to nucleosomes only in the presence of HMGB1. B. Binding profile of ER binding to nucleosomes in the presence of 400 nM HMGB1. Phosphoimager intensities of DNA and complex band were determined and % complex plotted using the Origin 6.2 software. ER concentrations were 0, 10, 20, 30, 40, 50, 60 & 80 nM. The profile for ER binding to nucleosome without HMGB1 could not be plotted because ER binding is not significant without

HMGB1. 199

100 B 80

% Complex 20

0 110 ER (nM)

200

Appendix 3. ER binding to 3E1 nucleosomes in the presence of 400 nM HMGB1.

Nucleosomes were incubated for 30 mins with increasing concentration of ER on ice, and run on 4% polyacrylamide gel for 2 hours. Lanes 1-8 represents ER concentrations 0, 10,

20, 40, 50, 60 & 80 nM, respectively. ER binds significantly to nucleosomes only in the presence of HMGB1. B. Binding profile of ER binding to nucleosomes in the presence of

400 nM HMGB1. Phosphoimager intensities of DNA and complex band were determined and % complex plotted using the Origin 6.2 software. ER concentrations were

0, 10, 20, 30, 40, 50, 60 & 80 nM. The profile for ER binding to nucleosome without

HMGB1 could not be plotted because ER binding is not significant without HMGB1.

201

B 80

% Complex 20

110 ER (nM)

202

Appendix 4. ER binding to 4E0 nucleosomes in the presence of 400 nM HMGB1.

Nucleosomes were incubated for 30 mins with increasing concentration of ER on ice, and run on 4% polyacrylamide gel for 2 hours. Lanes 1-8 represents ER concentrations 0, 10,

20, 30, 40, 50, 60 & 80 nM, respectively. ER binds significantly to nucleosomes only in the presence of HMGB1. B. Binding profile of ER binding to 4E0 nucleosomes in the presence of 400 nM HMGB1. Phosphoimager intensities of the DNA and complex bands were determined and % complex plotted from three independent gels, using the Origin

6.2 software. ER concentrations were 0, 10, 20, 30, 40, 50, 60 & 80 nM. The profile for

ER binding to nucleosome without HMGB1 could not be plotted because ER binding is not significant without HMGB1.